- Email: [email protected]
Integration of Endocrine Signals That Regulate Insect Ecdysis
Frontiers in Neuroendocrinology 23, 179 –199 (2002) doi:10.1006/frne.2002.0228, available online at http://www.idealibrary.com on
Integration of Endocrine Signals That Regulate Insect Ecdysis Karen A. Mesce* and Susan E. Fahrbach† *Departments of Entomology and Neuroscience and Graduate Program in Neuroscience, University of Minnesota, St. Paul, Minnesota 55108; and †Department of Entomology and Neuroscience Program, University of Illinois at Urbana–Champaign, Urbana, Illinois 61801
The extremely large number of insects and members of allied groups alive today suggests that molting—shedding of an old cuticle—may be one of the most commonly performed behaviors on our planet. Removal of an old cuticle in insects is associated with stereotyped, species-specific patterns of behavior referred to as ecdysis. It has been recognized for decades that the initiation of ecdysis is under hormonal control, but until recently many of the key peptides that regulate ecdysis were unknown. The report in 1996 of a new ecdysis-triggering hormone (ETH) sparked an era of significant advances in our understanding of the regulation of molting. This article summarizes the current model of peptide regulation of ecdysis, a model that is based on a positive feedback loop between ETH and a brain peptide, eclosion hormone. Then the relationship of these regulatory peptides to the neural circuitry that is the ultimate driver of the behavior are described. Because insects can undergo both status quo (larval–larval) and metamorphic (larval–pupal and pupal–adult) molts, differences in ecdysis behavior at different life stages are described and potential sources of these differences are identified. Most of the work described is based on studies of ecdysis in the hawkmoth, Manduca sexta, but results from studies of ecdysis in the fruit fly Drosophila melanogaster are also discussed. KEY WORDS: bursicon; CCAP; ecdysis; ecdysis-triggering hormone; eclosion; eclosion hormone; epitracheal glands; Inka cell; Manduca sexta; molting; VM neuron. ©
2002 Elsevier Science (USA)
INTRODUCTION
Insect species appear in the fossil record approximately 425 million years ago and today abound by the millions, a testimonial to their differential reproductive success. In part, this success, which reflects an extraordinary ability to adapt to varying environmental conditions, can be attributed to their sophisticated endocrine systems. The rich behavioral repertoires expressed by insects owe their flexibility and operational range to an overlay of endocrine factors that expands the capabilities of the relatively simple arthropod nervous system. All aspects of an insect’s life are intimately associated with its endocrinology. Chief among the insect hormones are the ecdysteroids (steroids), juvenile Address correspondence and reprint requests to S. E. Fahrbach, Department of Entomology, University of Illinois at Urbana–Champaign, 320 Morrill Hall, 505 S. Goodwin Avenue, Urbana, IL 61801. Fax: (217) 244-3499. E-mail: [email protected]. 179 ©
0091-3022/02 $35.00 2002 Elsevier Science (USA) All rights reserved.
180
MESCE AND FAHRBACH
hormones (terpinoids), peptides, and biogenic amines (55). Recent reviews have examined the actions of neuropeptides on visceral tissues, muscle, and the central and peripheral nervous systems (4, 53, 60, 63). The biological functions of octopamine, a key neuromodulator-neurohormone in insects, have also been recently reviewed (3) and therefore are not considered here. Other recent reviews have considered the ecdysteroids as mediators of neural plasticity (19) and provide an historical perspective on our understanding of the neuropeptide regulation of ecdysis behavior (14). The present review has more narrow goals. One is to describe how a remarkably elaborate cascade of endocrine factors regulates the seemingly simple behavioral task of shedding the cuticle in a way we believe will be accessible to the nonspecialist. A second goal is to compare what is known about the control of ecdysis behavior in larvae, pupae, and adults, the three major life stages of insects with complete metamorphosis. Our aim is to integrate a series of recent studies and unpublished results to illustrate the intricate neuroendocrine relationships required for expression of ecdysis-related behaviors. As one might expect, the study of insect molting requires investigation at both organismal and cellular–molecular levels. Physiological and behavioral studies have traditionally used large insects to study ecdysis, particularly the tobacco hornworm Manduca sexta (18, 70, 76). Such studies are now profitably being extended with molecular genetic studies of the fruit fly Drosophila melanogaster, the first postgenomic insect. Because progress in our understanding of the hormonal regulation of ecdysis cascades is currently rapid, this review augments other recent reviews on the topic (25, 29, 52, 73).
HISTORICAL PERSPECTIVE AND OVERVIEW OF ECDYSIS
This is still an era of discovery for insect hormones. The number of hormones known to be involved in the regulation of ecdysis-related motor programs has grown significantly over the past 5 years. Because the accompanying nomenclature is potentially confusing, careful attention is given to the definition of key terms used in this article. At the end of each molt cycle (the production of a new cuticle by the monolayer of epidermal cells that produce the insect exoskeleton), insects perform stereotyped, species-specific behaviors to shed the cuticle of the previous stage in a process termed “ecdysis”. A separate term, eclosion, is typically reserved for the final ecdysis of the insect’s life, so that adult ecdysis is often referred to as adult eclosion or simply eclosion. Over 30 years ago, blood-borne eclosion hormone (EH) 1 activity was first defined on the basis of its ability to 1 Abbreviations used: 20-E, 20-hydroxyecdysone; A1, A2 etc., abdominal ganglia of the ventral nerve cord; CCAP, crustacean cardioactive peptide; cGMP, cyclic 3⬘,5⬘-guanosine monophosphate; CNS, central nervous system; DL, dorsal longitudinal muscle; EH, eclosion hormone; ETH, ecdysis-triggering hormone; LDI, long distance interneuron; MN-2, motor neuron 2 of Manduca abdominal ganglia; MN-3. motor neuron 3 of Manduca abdominal ganglia; PETH, pre-ecdysis-
HORMONES AND INSECT MOLTING
181
initiate ecdysis behavior in pharate adult giant silkmoths (70, 76). (The term “pharate” refers to an insect fully developed and ready to molt, but still encased in the cuticle of the preceding stage.) Subsequent studies in the now more commonly used North American hawkmoth, Manduca sexta (hereafter referred to as Manduca), have revealed that EH also elicits ecdysis behavior when the insect molts from one larval stage to the next (termed larval ecdysis) and from the larval to the pupal stage (pupal ecdysis). Eclosion hormone is now identified as a 62-amino-acid peptide synthesized exclusively in two pairs of ventral medial (VM) neurons located in the protocerebral region of the brain (26, 32–34, 45, 54, 75). Although Manduca is the primary species used for research into the effects of EH on behavior, EH has also been shown to be synthesized by VM neurons and to initiate ecdysis behavior in nonlepidopteran insect species, including Drosophila (30, 73, 77). Eclosion hormone is currently assumed to regulate all insect ecdyses. The central nervous system (CNS) of insects consists of a brain and a chain of segmental neural ganglia, referred to as the ventral nerve cord (5). The motor neurons that drive the motor patterns of ecdysis reside in the segmental ganglia of the abdomen and the thorax; their axons innervate the musculature of the exoskeleton via segmentally repeated dorsal and ventral nerves (42). The life cycle of Manduca consists of five larval stages, a pupal stage during which adult development occurs, and the adult (the life cycle of Drosophila is similar, although there are only three larval stages in this species.) Each stage is separated by an episode of ecdysis in which the cuticle of the former stage is shed. In the case of Manduca and many other insects, the performance of ecdysis behavior removes the old cuticle by a series of coordinated abdominal movements characterized by an element of an anteriorly directed peristalsis. Adult eclosion in Manduca, however, differs from both larval and pupal ecdysis in one highly significant aspect: No consistent pattern of preecdysis rhythmic motor activity is expressed as a prelude to adult eclosion. In sharp contrast to pharate larvae and pupae, the pharate adult remains fairly quiescent except for occasional bouts of body twitching (Fig. 1A). Prior to each molt, hemolymph titers of ecdysteroids (primarily 20-hydroxyecdysone, or 20-E) rise and then return to basal levels (55); without the decline, ecdysis is blocked, and recent studies indicate that both central and peripheral actions of the steroid are important in preparing the insect for each molt (35, 84, 85). The decline in circulating steroids likely acts in concert with other signals that either specify the insect’s readiness to molt or provide circadian cues so that the insect molts during the appropriate phase of the light– dark cycle (74).
triggering hormone; SEG, subesophageal ganglion; T1-T3, the prothoracic, mesothoracic, and metathoracic ganglia of the ventral nerve cord; TSM, tergosternal muscle; TX, thorax; VM, ventral medial neurons of the insect brain that synthesize and secrete EH.
182
MESCE AND FAHRBACH
FIG. 1. Extracellular recordings from the tergosternal muscles (TSMs) in the fourth through sixth abdominal segments (A4 –A6) and from the dorsal longitudinal (DL) muscles in the thorax (TX) of an intact pharate adult Manduca. Muscle activity was monitored beginning approximately 5– 6 h before the predicted time of adult ecdysis to determine if any pre-ecdysis-like motor patterns were present. Electrophysiological methods used are as previously described (49). (A) Records of muscle activity expressed throughout various time periods prior to eclosion are shown; samples of activity patterns presented were randomly chosen. In over 40 insects examined in this way, no preecdysis motor pattern was observed. Activity in the thorax was minimal and remained unpatterned until eclosion. Intersegmental TSM coactivity was always variable, of short duration (reflecting a rapid body twitching), and intermittent. (B) At the onset of eclosion behavior, the DL muscles first showed strong activity, which was always coactive with TSM excitation and correlated with synchronous lateral body contractions that caused the abdomen to extend. By the time of the sixth extension cycle, the peristalsis component appeared (marked by asterisks) and continued, while involving more posterior segments (not shown). This rapid anteriorly directed peristalsis caused the retraction of the abdomen, which was followed by the abdominal extension, a pattern that was repeated for many cycles. The temporal parameters of these extension bouts were indistinguishable from the activity patterns expressed during larval and pupal preecdysis behavior.
HORMONES AND INSECT MOLTING
183
FIG. 2. (A) Schematic diagram of the peripheral peritracheal neuroendocrine system of insects. (B) Summary diagram of the hormonal and cellular interactions leading to events involved in ecdysis-related behaviors derived from research on Manduca and Drosophila. Boxes contain both the names of ecdysis-related hormones (or factors) and the specific cells that secrete them. Hatched lines indicate relationships that are suggested by current research and for which comparatively less is known.
PREECDYSIS BEHAVIOR AND ITS ENDOCRINE CONTROL
The distinctive preecdysis motor patterns observed in larval and prepupal insects loosen the attachments between the new and the old cuticle prior to ecdysis. In Manduca larvae and prepupae, two distinct phases of preecdysis behavior, designated preecdysis I and preecdysis II, have been identified (84). These preecdysis behaviors are regulated by peptide hormones secreted by 18 segmentally repeated epitracheal glands located near the spiracular tracheae of the abdominal segments (Fig. 2A). The epitracheal glands, which were first described in 1996 (37, 83), each consist of a small number of cells (four per gland in Manduca), one of which is referred to as the Inka cell. Inka cells are characterized by synthesis and secretion of a myomodulin-like peptide (59) and
184
MESCE AND FAHRBACH
at least three additional peptides, two of which, preecdysis-triggering hormone (PETH) and ecdysis-triggering hormone (ETH), activate preecdysis I and preecdysis II (83, 84). The myomodulin-like peptide is released from the Inka cells of Drosophila and Manduca in the hour prior to all ecdyses and is present in putative Inka cell homologs in various insects such as beetle, dragonfly, and mosquito, but at present nothing is reported concerning its ecdysis-related functions (59). PETH and ETH were first described in Manduca and have been subsequently identified in another lepidopteran, the silkmoth, Bombyx mori (1), and in Drosophila (61). Both peptides are encoded by a single gene downstream of a promoter region that contains ecdysone response elements (84). In the Inka cells, a precursor peptide is cleaved to produce the 11-amino-acid PETH, the 26-amino-acid ETH, and a 46-amino-acid peptide of unknown function referred to as ETH-associated peptide (ETH-AP). Application of PETH to the isolated CNS of Manduca can activate a known neural correlate of preecdysis I behavior, strong bursts of motor activity in the dorsal nerves. This pattern of electrical activity in the CNS is often referred to as the fictive component of preecdysis behavior. The bursts in the dorsal nerves are in phase with the synchronous dorsoventral compressions of the insect’s abdomen that occur during the first 20 or so min of preecdysis behavior (51, 56, 57, 84). Animals injected only with PETH express preecdysis I behavior exclusively, and no other behaviors associated with the ecdysis sequence such as preecdysis II or ecdysis are subsequently displayed (84). In contrast to the actions of PETH, currently believed to be restricted to controlling preecdysis, ETH also indirectly activates the behavioral routines of ecdysis itself in a chain of events that are described in more detail in following sections (83). The most direct action of this peptide, however, is the initiation of preecdysis II behavior (84). Injections of low physiological concentrations of ETH into pharate larvae previously exposed to PETH result in the selective expression of asynchronous posterior-ventral contractions of abdominal body segments coupled with repeated retraction-extensions of the prolegs. A fictive behavioral correlate of this preecdysis II pattern has also been identified: it consists of strong motor bursts in the ventral nerves in the absence of strong bursting in the dorsal roots (84). Fictive preecdysis II can be elicited with low concentrations of ETH. Application of higher ETH concentrations (100 –300 nM) to isolated ventral nerve cord preparations can elicit ecdysis behavior, with latencies in the range of 22–30 min (84). Under natural conditions, preecdysis I, preecdysis II, and ecdysis occur in sequence. The strong dorsoventral contractions of preecdysis I (roughly a 1-s contraction followed by 3 s of relaxation) are observed first. Preecdysis II behavior in Manduca is then typically observed after the first third of preecdysis I behavior has been performed and continues along with expression of the preecdysis I program until ecdysis is initiated, about 40 min later [see Fig. 6B in reference (84)]. PETH and ETH appear to be co-released from the Inka cells, but a lower response threshold to PETH ensures that preecdysis I is expressed first followed by preecdysis II and then by ecdysis (82, 84).
HORMONES AND INSECT MOLTING
185
ECDYSIS BEHAVIOR AND ITS ENDOCRINE CONTROL
Although the names ascribed to EH and ETH imply that they are the direct hormonal activators of the neural circuits responsible for ecdysis behavior, to date there is no evidence that either hormone is tied to such proximal activation. Instead, it is the neuropeptide crustacean cardioactive peptide (CCAP) that has been shown to have the most direct, shortest latency route to activation of the ecdysis motor program (22). Several studies have indicated that EH stimulates the release of CCAP from neurosecretory neurons distributed along the ventral nerve cord (23, 66). In Manduca, this network consists primarily of segmentally repeated lateral pairs of neurons named cell 27 [or NS-L(neuromere no.)] and interneuron/cell (IN)-704 (7, 12, 38), often referred to jointly as 27/704. These cells were initially identified on the basis of their immunoreactivity to an antibody prepared against CCAP from crustaceans (67) and hence are referred to as CCAP-ir cells. Several lines of evidence indicate that authentic CCAP is present in the insect nervous system. A nonapeptide identical in amino acid sequence to that of CCAP originally isolated from a shore crab, Carcinus maenas, was purified from extracts of the Manduca CNS (6, 41). More recently, the CCAP gene of Manduca was cloned, and it was demonstrated using in situ hybridization that almost all CCAP-ir cells in the CNS express the mRNA coding for this peptide (43). The distribution of CCAP neurons is highly conserved between insects and crustaceans (62, 69). Although extant insect species and crustaccans have evolved as independent lineages for hundreds of millions of years (20, 24, 78), CCAP has been recently shown to be released during the ecdysis of crabs and crayfish (62), suggesting that its behavior-activating functions have also been conserved. Intracellular recordings made from abdominal CCAP neurons in pharate larvae of Manduca revealed that EH rapidly elevated cGMP levels within these cells, which in turn resulted in increased release of CCAP (23). When latencies to ecdysis were compared after application of various peptides to the isolated nervous system, the response to CCAP (5 min) was significantly shorter than the response to either EH (26 min) or ETH (40 min) (23). CCAP can also induce ecdysis at developmentally inappropriate times (i.e., intermolt stages, when the new cuticle is incomplete), periods when the CNS is refractory to ETH and EH action (23). At present, therefore, no peptide is linked more tightly to the onset of ecdysis than CCAP. This makes it particularly unfortunate that no information is currently available about the neuronal targets of CCAP at the level of ecdysis pattern-generating circuitry. Although a number of motor neurons with ecdysis-related patterns of activity have been identified (79), the individual central pattern generator neurons that generate the motor patterns underlying ecdysis behavior have not been identified. This is clearly a challenging area for future research. Figure 2B shows the coordinated hormonal events associated with the expression of ecdysis behavior. To summarize, at the end of the molt cycle, when the new cuticle is formed and a successful molt is possible, the Inka cells of the
186
MESCE AND FAHRBACH
epitracheal glands and the VM neurons of the brain release their respective peptide hormones, PETH/ETH and EH (27, 35, 84). At present it cannot be conclusively stated that either EH or ETH is released first, as each hormone has been clearly demonstrated to cause the release of the other (13, 23, 36). Arguments have been made for the primacy of PETH/ETH based on in vitro differences in the responses to PETH, ETH, and EH (82). What is clear is that reciprocal positive feedback mechanisms ensure the attainment of successively higher levels of PETH/ETH and EH. When sufficiently high levels of circulating ETH are achieved, the VM neurons of the brain respond by producing a surge of EH released both centrally and peripherally into the hemolymph. Such a coordinated release is made possible by the extensive projections of the VM neurons within each segmental ganglion as well as projections to a peripheral neurohemal release site via the proctodeal nerve (75). One of the actions of EH on the CCAP-containing 27/704 cells is to facilitate the release of CCAP (23). CCAP then serves as the proximate trigger of ecdysis behavior.
BURSICON IN THE CRUSTACEAN CARDIOACTIVE PEPTIDE NETWORK
Adding a further level of complexity, a subset of the neurons of the CCAP network also contain a neuropeptide called bursicon. Bursicon was originally defined as the primary factor that promotes the hardening and tanning (sclerotization) of new cuticle after the insect has completed ecdysis (64). New studies indicate that, at a minimum, the CCAP cell 27 in Manduca and its homologs in other insects likely contain bursicon (39, 28). An attempt to determine the complete amino acid sequence of the bursicon hormone is currently underway in the laboratory of H.-W. Honegger (Vanderbilt University, TN). The microsequencing of bursicon hormone purified from cockroaches recently yielded several partial peptide sequences (40). Subsequent immunocytochemical studies using antisera directed against these partial sequences confirmed that the CCAP-ir cell 27 in Manduca and the CCAP-ir cell 27 homologs in other insects, including crickets, cockroaches, and several species of flies, are immunoreactive to these antisera (28). These results suggest that specific bursicon amino acid sequences are conserved even among distantly related insect species. Studies in Manduca have demonstrated an association between EH release and the subsequent secretion of bursicon into the hemolymph, although a full picture is lacking because published reports have focused almost exclusively on adult eclosion (64, 71). The important issue of how the actions of bursicon are temporally restricted, however, remains unresolved. For the Manduca pharate adult, in particular, it is essential that tanning of the cuticle not be activated until the insect has removed its old cuticle, surfaced from its underground pupation site and gained a perch from which it can inflate its wings. The premature action of bursicon before completion of the ecdysis motor program could result in a premature hardening of the cuticle and a blocking of wing inflation that would likely be lethal. The observation that CCAP and
HORMONES AND INSECT MOLTING
187
bursicon are often colocalized in cell 27 therefore provides a neuroendocrine puzzle. Studies are needed to determine whether these hormones are differentially secreted. Whether bursicon is modified in some way to produce its delayed (45– 60 min) action during adult ecdysis or has additional actions that block premature sclerotization are questions that remain unanswered.
ADDITIONAL FACTORS CONTROLLING ECDYSIS
Although strong evidence indicates that EH is the link between ETH and CCAP action (23), a number of intriguing recent studies suggest the presence of additional layers of control, some of which involve descending neural activity (2, 58, 82). Ecdysis-triggering hormone has been shown to excite the EH-containing VM neurons (13) and to elicit a broadening of the VM cell spike (23). Electrical stimulation of these cells causes elevation of cGMP in the CCAP 27/704 neural network. However, when the part of the brain (protocerebrum) that contains the VM neurons was removed from pharate Manduca larvae (the deutocerebrum was also removed, leaving the tritocerebral lobes intact), 10 of 14 insects (71%) injected with ETH expressed ecdysis behavior at the expected time (82). Removal of the subesophageal ganglion (SEG) or thoracic ganglia severely limited or prevented the onset of ecdysis behavior. In turn, application of ETH to isolated nerve cords with the SEG intact but the brain ablated elicited an unmistakable ecdysis motor response and concomitant elevations in cGMP among the CCAP cells. Such results suggest that, although ETH-induced release of EH from the VM neurons is sufficient to activate signaling cascades resulting in CCAP release, this pathway may not be required. Studies in Drosophila (2, 46) further suggest that the VM neurons in the brain may not be required for ecdysis. Genetically modified knockout flies that lack the VM neurons expressed components of adult eclosion behavior and emerged from the pupal cuticle; the behavior of these pharate adults, however, was described as weak and poorly organized. In addition, the flies that did succeed in eclosing could not inflate their wings, possibly as a consequence of abnormal bursicon release (46). In addition, the pharate adult knockout flies lacking the VM neurons were no longer sensitive to ETH. Although ETH has not yet been shown to activate a specific behavior in pharate adult flies, treatment of pharate adult flies with exogenous ETH can advance the onset of adult eclosion (2, 46), and a gene encoding ETH-like peptides has been identified in Drosophila (61). The lack of a response to ETH in the knockout flies suggests that, in wild-type flies, ETH, EH, and/or sensory inputs must converge on some unidentified neural or hormonal element(s) in the CNS in order to drive activity in the CCAP cell 27/704 network. In the absence of the ETH-sensitive VM neurons, ecdysis activation can apparently still be achieved by direct activation of downstream parts of the pathway. Whether these observations on Drosophila can be generalized to other insects is currently unknown.
188
MESCE AND FAHRBACH
In addition, the switch from larval preecdysis to ecdysis behaviors appears not to be a simple consequence of CCAP release. Studies by Zitnan and Adams (82) have demonstrated that an inhibitory influence from the SEG, thoracic ganglia, and first abdominal ganglion must be removed before CCAP-induced ecdysis behavior can be initiated. After initiation of spontaneous or ETHevoked preecdysis behavior, isolation of the abdomen from anterior ganglia (ligation) significantly advanced the onset of ecdysis in all insect tested. Removal of the head has also been shown to induce eclosion onset in pharate adult flies and in Manduca (13). Because ligations that separate the head from the body in pharate adults can elicit eclosion within minutes or even seconds (K. A. Mesce, personal observation), such massive severing of neural connections may result in an injury-induced excitation of stimulatory descending neural signals rather than a simple removal of inhibitory elements. Such a scenario is typically overlooked. A picture emerges in which descending neural control mechanisms are coupled with hormonal feedback loops to ensure that an insect initiates ecdysis only when the insect is physiologically prepared to survive the ecdysis event (the formation of the new cuticle is complete and the digestion of the old cuticle is sufficiently advanced). As previously noted, which trigger has primacy is currently impossible to say and may differ among insect species.
CRUSTACEAN CARDIOACTIVE PEPTIDE NEURONS AND STAGE-SPECIFIC BEHAVIORS
The morphology of the CCAP cells indicates that both central and peripheral release are possible (7, 15, 38). It has been argued that the CCAP-containing neurons active at the time of ecdysis are those that show immunodetectable elevations in cGMP (12, 21). Although other CCAP neurons are present in the CNS, only the cell 27/704 network, consisting of approximately 50 neurons, expresses cGMP at the time of ecdysis. During adult and pupal ecdysis, however, only the CCAP-ir 27s and not the 704s in the abdomen express increased cGMP (16), while at larval ecdysis the abdominal 704s also express increased cGMP. Thus, local spatial differences in the release of CCAP could account for differences in the activation of ecdysis-related motor programs displayed by different life stages. It was originally noted that the entire cell 27/704 network becomes cGMPimmunopositive about the same time. One recent study, however, has shown that in untreated pharate pupae the 27/704 cells of the SEG and thoracic ganglia show weaker but still detectable cGMP immunoreactivity at 6 h before the onset of ecdysis, well before the release of PETH and ETH from the Inka cells. Zitnan and Adams (82) have suggested that this 27/704 cGMP-ir network in the thoracic ganglia could release an hypothetical ecdysiostatic factor responsible for the descending inhibitory influences described in the previous section. These authors, however, based their idea on the misconception that the thoracic 27/704 network in larvae does not contain CCAP, therefore lead-
HORMONES AND INSECT MOLTING
189
ing them to suggest that this network possesses a different neurohormone. Although there are indeed differences among the thoracic and abdominal 27/704 neurons, as described below, the lack of CCAP is not one of them. Confusion over CCAP expression in the thoracic ganglia stems from several sets of independently published studies in which the results of CCAP immunostaining differed. The study by Klukas et al. (38; see their Figs. 8 and 9) found that the SEG 27/704 cells and the thoracic cell 27s are CCAP-ir in larvae and developing adults. By contrast, another study reported no CCAP staining after the larval stages (8), and other authors mentioned but did not show that the thoracic homologs always express at least weak CCAP immunoreactivity (13). The recent identification of the gene for CCAP in Manduca provides clarification of this issue (43). The published report indicates that there is likely only one CCAP gene in this species. The spatial expression pattern of the CCAP gene using in situ hybridization techniques confirms that the CCAP-ir cells in the larval SEG and thoracic ganglia indeed express the CCAP gene. Later, in the developing adult, the thoracic CCAP-ir 27 and 704 cells continue to exhibit robust staining (Fig. 3). A striking observation that indicates the strength of the association of this peptidergic neural network in Manduca with ecdysis is that, after the final performance of ecdysis by the pharate adult, the CCAP network, with the exception of one cell pair, is dismantled by the programmed cell death of the CCAP neurons (17). Curiously, CCAP expression in the thoracic 27/704 cells of Manduca is highly variable even during larval life as compared to that in the abdominal ganglia. This is especially evident in pharate fifth instar larvae. It is unknown whether such variability represents the transient inability of thoracic cells to generate transcript or reflects intermittent cycles of CCAP synthesis. Other intriguing differences between the thoracic and abdominal homologs have also been documented (15, 38). For example, a gentler tissue fixation protocol is required to immunostain the 27/704 cells of the thoracic ganglia with CCAP antiserum, as if the CCAP epitope is more difficult to preserve in this region of the central nervous system. Differences in CCAP fixation requirements may point to underlying differences in the thoracic cells with regard to CCAP content, storage, and release. Also notable are the other developmentally regulated epitopes and proteins that are differentially expressed among the 27/704 thoracic and abdominal homologs. In fifth instar larvae of Manduca, for example, the 27 cells in the SEG and the 27/704 cells in the three thoracic ganglia all express a 27-kDA membrane-associated protein (38); subsequent microsequencing has revealed novel partial amino acid sequences (K. A. Mesce, unpublished observations). At no time during development do any cell types in the CNS other than the 27/704s express this antigen. A similar pattern of antigen expression in the thoracic ganglia was also evident in larvae stained with the monoclonal antibody P2C5 (17). Studies of P2C5 expression revealed that the 27 cells in the abdomen became P2C5-ir during the pupal and adult stages of development, while comparable cells in the thorax were immunopositive at all life stages. Elimination of 20-E fluctuations during the larval– pupal transition by ligation prevented expression of the P2C5 antigen in these
190
MESCE AND FAHRBACH
FIG. 3. The 27/704 neurons (arrow) immunostained with an antiserum against crustacean cardioactive peptide (CCAP) and a secondary antibody conjugated to Cy3. CCAP-immunoreactive neurons were observed in all ganglia, including the prothoracic ganglion (shown here), of a developing adult 2 days prior to eclosion (Day: ⫺2). Immunostaining protocols and antiserum are as previously described (17, 38). Robust CCAP staining of the 27/704 cells was present in pharate adults and then declined after eclosion, when these cells underwent programmed cell death (17). Scale bar ⫽ 100 m.
abdominal cells (S. E. Fahrbach and C.-M. Wang, unpublished observations), suggesting that regulation of the P2C5 antigen is under the influence of this steroid. Future investigations into the identities and functions of the cellular proteins and antigens just discussed may provide useful information about why and how the thoracic CCAP-27/704 cells differ from those in the abdomen. Because both antigens are associated with cells that express cGMP in response to ETH and EH release, the possibility cannot be dismissed that these factors represent hormone receptor elements and associated signaling proteins.
EXPRESSION OF NOVEL MOTOR PATTERNS AT ADULT ECDYSIS
We now wish to address directly the mechanisms that result in the dramatic differences between the ecdysis-related behaviors of the pharate adult Manduca and those of pharate larvae and pupae (Fig 1B). As stated, the most
HORMONES AND INSECT MOLTING
191
conspicuous difference is the absence of preecdysis I and preecdysis II behaviors prior to the expression of ecdysis behavior (Fig. 1A). Instead of loosening its cuticle via a preecdysis motor program, the pharate adult relies on a series of rotary abdominal movements that are displayed about 10 –12 h before ecdysis. This preparatory behavior, so named to distinguish it from the preecdysis behavior of the larva and the prepupa, is apparently not dependent on ETH or PETH, but instead depends on the decline in ecdysteroid titers that precedes ecdysis (72). Such observations raise several questions. Are preecdysis neural circuits lost during the adult development that occurs after pupal ecdysis? What roles do PETH and ETH play during the last ecdysis? One hypothesis is that the preecdysis pattern generator is retained into the pharate adult stage, but is used to create a novel motor component specific to the adult ecdysis rhythm. The essence of this idea is that separate preecdysis and ecdysis neural circuits are activated together to form a conjoint rhythm, in much the same way as has been demonstrated for neural circuits underlying stomatogastric rhythms in crustaceans (9 –11, 44, 68). The pharate adult performs a repeated series of abdominal extensions and retractions that rely, in part, on the activity of MN-2 and MN-3, the same motor neurons activated during preecdysis and ecdysis behaviors in larvae and pupae. The abdominal retraction phase in the adult consists of a rapidly progressing peristalsis that is likely generated by conserved larval–pupal ecdysis circuitry (49). Though not expressed prior to adult ecdysis, if the preecdysis I neural circuit were operational at the time of ecdysis the motor output of this circuit could constitute the rhythmic extension component observed during adult ecdysis. Support for this idea comes from the analysis of MN-2/MN-3 motor bursts that are repeatedly generated during the extension phase of adult ecdysis. This analysis was accomplished by recording from the tergosternal muscles (TSMs) innervated exclusively by the identified motor neurons MN-2/MN-3, cells used earlier in development to generate preecdysis and ecdysis patterns by larvae and pupae (K. A. Mesce, unpublished observations). Remarkably, the temporal aspects of MN-2/MN-3 (TSM) extension bursts are indistinguishable from MN-2/MN-3 bursting activity generated by larvae during preecdysis behavior. Based on the data of Miles and Weeks (50) for preecdysis activity, the mean period between successive MN-2/MN-3 preecdysis bursts was 8.4 ⫾ 1.2 s (N ⫽ 5), not statistically different from the adult TSM bursts (8.24 ⫾ 0.95 s, N ⫽ 5; p ⬍ 0.001, two-tailed t test). The duration of adult TSM bursts was also similar to the duration of MN-2/MN-3 preecdysis bursts (range of 1.6 –5s). In over 40 animals examined, ecdysis behavior was always marked by three to eight cycles of MN-2/MN-3 coactivity in the absence of the rapid peristaltic (retraction) component. This rapid peristalsis eventually emerged and became interspersed between the extension bursts (Fig. 1B). The period between successive TSM extension bursts remained stable throughout eclosion (49). Independent evidence indicates that an artificial exposure of the isolated CNS to ETH and CCAP can result in the simultaneous expression of ecdysis and preecdysis motor bursts (22). Thus, the novel ecdysis behavior expressed by the pharate adult may be a relatively simple consequence of a conserved ETH-
192
MESCE AND FAHRBACH
activated preecdysis circuit activated simultaneously with the CCAP-dependent circuit responsible for ecdysis. One last point is that the original muscle targets of MN-2/MN-3 (tergopleural and anterior lateral external muscles) are replaced during adult development with the TSMs, which consist of a larger mat of dorsoventrally oriented muscles. Thus, although the magnitude of lateral body constrictions is greater in the adult than in the larva or prepupae, this is not necessarily a consequence of motor neuronal rewiring but could rather reflect muscle remodeling during development. The temporal coordination and modulation of the preecdysis and ecdysis circuits are likely dependent on descending neural inputs, as it has been shown that the removal of descending activity below A2 causes an immediate switch from the adult-specific ecdysis motor pattern to that of the larval-like one (49). Two segmental homologs have been identified that project uninterrupted from A1 and A2 to the terminal abdominal ganglion (Fig. 3). These long distance interneurons (LDIs), when electrically stimulated, cause coactivity of the abdominal TSMs similar to that observed during larval or pupal preecdysis behavior and during the extension phase of adult ecdysis. Because little is known about the CCAP and ETH neuronal targets responsible for ecdysis pattern generation, these newly identified LDI cells provide a potentially fruitful area for future studies of hormonal targets underlying rhythmic behaviors. In addition, because these LDI neurons are present in larvae (K. A. Mesce, unpublished observations), the LDIs could contribute to the descending neural inhibition of ecdysis that can be removed when the connectives are severed between A1 and A2 (82). Interestingly, when larval abdominal ganglia already exposed to CCAP remained connected to T1 and A1, abdominal ganglia 4 – 6 expressed mixed preecdysis and ecdysis bursts. Only when the abdominal ganglia were isolated from T3 and A1 did all of the remaining ganglia exhibit a strong and pure ecdysis motor rhythm. For comparison in the adult, removal of A1 (and, by necessity, A2, as both are part of a larger fused pterothoracic ganglion in the adult) elicits similar strong, larval-like ecdysis bursts. In the case of the adult such activity reflects the unmasking of a motor pattern never normally expressed in its pure larval-like form. In summary, even after EH and CCAP have been released, descending inhibitory neural influences may coordinate the precise timing of ecdysis initiation at all developmental stages. Specifically, in larvae and pupae, the sequential temporal coordination of preecdysis and ecdysis appears to be influenced by descending activity. In the adult, however, possibly the same descending influences may promote a coactivation of the preecdysis and ecdysis motor programs, resulting in the novel ecdysis behavior of the adult (Fig. 4).
CIRCADIAN REGULATION OF ECLOSION: HORMONAL AND CLOCK-OUTPUT MECHANISMS OF BEHAVIOR
One additional important difference between larval–pupal and adult ecdysis behavior in Manduca and Drosophila is that adult eclosion behavior is under
HORMONES AND INSECT MOLTING
193
FIG. 4. Descending long distance interneurons (LDIs) filled retrogradely with the tracer Neurobiotin. Tissue was subsequently reacted with a Cy5-conjugated streptavidin and imaged with the confocal microscope as previously described (48). Insects used were 2 days away from adult ecdysis (Day: ⫺2). (A) Staining of two prominent neurons whose axons in the connectives were filled with tracer after axons were transected below abdominal ganglion 4 (A4). Stained neurons in the first and second thoracic ganglia (T1 and T2) were absent, but were present in the subesophageal ganglion (SEG) (not shown). Neuronal staining in this far anterior ganglion indicated that the lack of filled cells was not the inability of the tracer to reach the thoracic neuromeres. (B) Same Neurobiotin-filled LDIs (arrows) in A1 and A2 of the pterothoracic ganglion. These cells were filled at a level just anterior to the terminal abdominal ganglion (i.e., below A5). Neurons projecting from anterior ganglia to the terminal ganglion were stained in the brain, SEG (both not shown), A1, and A2, but not the thoracic ganglia. Scale bars ⫽ 100 m.
194
MESCE AND FAHRBACH
circadian control. Recent reviews (31, 80) have explored the multiple components and cell molecular mechanisms underlying the clock control of eclosion. Although EH release appears to be (65) under circadian control, recent studies support the idea that multiple clock pathways likely regulate the timing of eclosion, especially because insects lacking their EH-containing VM neurons still show a circadian regulation of eclosion (46). In Drosophila, a class of CCAP-ir neurons to which the cell 27 homologs belong contains the protein known as LARK, which is present in the cellular cytoplasm and shows diurnal changes in abundance (47, 81). This RNA-binding protein has been hypothesized to regulate a translation factor that may mediate CCAP synthesis, secretion, or perhaps neuronal excitability. What regulates LARK itself remains a mystery, but could involve the gated release of EH. Understanding clock mechanisms regulating CCAP content and release will surely provide a more complete picture of the neural and endocrine pathways regulating the expression of ecdysis across insect species One of many remaining puzzles is that the CCAP-ir homologs in Drosophila and other higher dipterans do not express an increase in cGMP at the time of ecdysis, although other cell types and neurons do express such EH-dependent cGMP increases during eclosion. Similarly, expression of the mIgG-like antigen, specific to the CCAP/cGMP-ir neurons of Manduca and other insects (discussed earlier in this article), has never been observed on the CCAP-ir neurons of Drosophila (K. A. Mesce, unpublished observation). Such results indicate that the cGMP-signaling pathways, so highly conserved in the majority of insect orders and through development, may have become modified during evolution of the cyclorrhaphous insects. This alteration likely relates to the unique forms of ecdysis behavior displayed by flies. Possibly, understanding the regulation of LARK may provide clues as to why the cGMP-signaling pathway is absent or reduced in amplitude in the CCAP cells of higher dipterans. FUTURE STUDIES
The diverse array of arthropod species and their disparate environmental constraints provide the comparative neuroendocrinologist with an opportunity both to understand how neuroendocrine control mechanisms become constructed through evolution and how they shift throughout the life of an individual. Yet to date our understanding is based on the study of a very small number of insect species. The greatest opportunities for insect neuroendocrinology in the 21st century will come from a broadening of our perspective beyond the limited number of species currently studied. As ever, shedding of old questions will result in the emergence of new ones for us to solve. ACKNOWLEDGMENTS The authors acknowledge the assistance of Kathleen Klukas in the preparation of the figures. J. Whitfield provided helpful comments on the phylogentic relationship of insects and crustaceans.
HORMONES AND INSECT MOLTING
195
REFERENCES 1.
Adams ME, Zitnan D. Identification of ecdysis-triggering hormone in the silkworm Bombyx mori. Biochem Biophys Res Comm 1997; 230: 188 –191.
2.
Baker JD, McNabb SL, Truman JW. The hormonal coordination of behavior and physiology at adult ecdysis in Drosophila melanogaster. J Exp Biol 1999; 202: 3037–3048.
3.
Braunig P, Pfluger HJ. The unpaired median neurons of insects. Adv Insect Physiol 2001; 28(28): 185–266.
4.
Broeck JV. Neuropeptides and their precursors in the fruitfly, Drosophila melanogaster. Peptides 2001; 22(2): 241–254.
5.
Burrows M. The Neurobiology of an Insect Brain. Oxford, UK: Oxford Univ. Press, 1996.
6.
Cheung CC, Loi PK, Sylwester AW, Lee TD, Tublitz NJ. Primary structure of a cardioactive neuropeptide from the tobacco hawkmoth, Manduca sexta. FEBS Lett 1992; 313: 165–168.
7.
Davis NT, Homberg U, Dircksen H, Levine RB. Crustacean cardioactive peptide immunoreactive neurons in the hawkmoth, Manduca sexta and changes in their immunoreactivity during postembryonic development. J Comp Neurol 1993; 338: 612– 627.
8.
Davis NT, Homberg U, Dircksen H, Levine RB, Hildebrand JG. Crustacean cardioactive peptide-immunoreactive neurons in the hawkmoth Manduca sexta and changes in their immunoreactivity during postembryonic development. J Comp Neurol 1993; 338(4): 612– 627.
9.
Dickinson PS. Interactions among neural networks for behavior. Curr Opin Neurobiol 1995; 5: 792–798.
10.
Dickinson PS, Fairfield WP, Hetling JR, Hauptman J. Neurotransmitter interactions in the stomatogastric system of the spiny lobster: One peptide alters the response of a central pattern generator to a second peptide. J Neurophysiol 1997; 77: 599 – 610.
11.
Dickinson PS, Mecsas C, Marder E. Neuropeptide fusion of two motor-pattern generator circuits. Nature 1990; 344: 155–158.
12.
Ewer J, De Vente J, Truman JW. Neuropeptide induction of cyclic GMP increases in the insect CNS: Resolution at the level of single identifiable neurons. J Neurosci 1994; 14(12): 7704 –7712.
13.
Ewer J, Gammie SC, Truman JW. Control of insect ecdysis by a positive-feedback endocrine system: Roles of eclosion hormone and ecdysis triggering hormone. J Exp Biol 1997; 200(5): 869 – 881.
14.
Ewer J, Reynolds SE. Neuropeptide control of molting in insects. In: Pfaff DW, Arnold A, Etgen A, Fahrbach SE, Rubin R, Eds. Hormones, Brain, and Behavior. San Diego, CA: Academic Press, in press.
15.
Ewer J, Truman JW. Increases in cyclic 3⬘,5⬘-guanosine monophosphate (cGMP) occur at ecdysis in an evolutionarily conserved crustacean cardioactive peptide-immunoreactive insect neuronal network. J Comp Neurol 1996; 370: 330 –341.
16.
Ewer J, Truman JW. Invariant association of ecdysis with increases in cyclic 3⬘,5⬘-guanosine monophosphate immunoreactivity in a small network of peptidergic neurons in the hornworm, Manduca sexta. J Comp Physiol A 1997; 181(4): 319 –330.
17.
Ewer J, Wang CM, Klukas KA, Mesce KA, Truman JW, Fahrbach SE. Programmed cell death of identified peptidergic neurons involved in ecdysis behavior in the moth, Manduca sexta. J Neurobiol 1998; 37: 265–280.
18.
Fahrbach SE. The regulation of neuronal death during insect metamorphosis. BioSci 1997; 47: 77– 85.
19.
Fahrbach SE, Weeks JC. Hormonal regulation of neural and behavioral plasticity in insects. In: Pfaff DW, Arnold A, Etgen A, Fahrbach SE, Rubin R, Eds. Hormones, Brain and Behavior. San Diego, CA: Academic Press, in press.
196
MESCE AND FAHRBACH
20.
Friedrich M, Tautz D. Ribosomal DNA phylogeny of the major extant arthropod classes and the evolution of myriapods. Nature 1995; 376: 165–167.
21.
Gammie SC, Truman JW. An endogenous elevation of cGMP increases the excitability of identified insect neurosecretory cells. J Comp Physiol A 1997; 180: 329 –337.
22.
Gammie SC, Truman JW. Neuropeptide hierarchies and the activation of sequential motor behaviors in the hawkmoth, Manduca sexta. J Neurosci 1997; 17: 4389 – 4397.
23.
Gammie SC, Truman JW. Eclosion hormone provides a link between ecdysis-triggering hormone and crustacean cardioactive peptide in the neuroendocrine cascade that controls ecdysis behavior. J Exp Biol 1999; 202: 343–352.
24.
Giribet G, Ribera C. A review of arthropod phylogeny: New data based on ribosomal DNA sequences and direct optimization. Cladistics 2000; 16: 204 –231.
25.
Hesterlee S, Morton DB. Insect physiology: The emerging story of ecdysis. Curr Biol 1996; 6: 648 – 650.
26.
Hewes RS, Truman JW. TYhe roles of central and peripheral eclosion hormone release in the control of ecdysis behavior in Manduca sexta. J Comp Physiol 1991; 168A: 697–707.
27.
Hewes RS, Truman JW. Steroid regulation of excitability in identified insect neurosecretory cells. J Neurosci 1994; 14(3/2): 1812–1819.
28.
Honegger H-W, Market D, Kostron B, Wilson M, Pierce L, Choi D, Mesce KA. Cellular localization of bursicon using antisera against a partial sequence of this insect cuticlesclerotizing neurohormone. J Comp Neurol, in press.
29.
Horodyski F. Neuroendocrine control of insect ecdysis by eclosion hormone. J Insect Physiol 1996; 42: 917–924.
30.
Horodyski FM, Ewer J, Riddiford LM, Truman JW. Isolation, characterization, and expression of the eclosion hormone gene of Drosophila melanogaster. Eur J Biochem 1993; 215: 221–228.
31.
Jackson FR, Schroeder AJ, Roberts MA, McNeil GP, Kume K, Akten B. Cellular and molecular mechanisms of circadian control in insects. J Insect Physiol 2001; 47: 833– 842.
32.
Kamito T, Tanaka H, Sato B, Nagasawa H, Suzuki A. Nucleotide sequence of cDNA for the eclosion hormnone of the silkworm, Bombyx mori, and the expression in a brain. Biochem Biophys Res Commun 1992; 31: 514 –519.
33.
Kataoka H, Li JP, Lui AS, Kramer SJ, Schooley DA. Complete structure of eclosion hormone of Manduca sexta. Assignment of disulfide bond location. Int J Pept Protein Res 1992; 39: 29 –35.
34.
Kataoka H, Troetschler RG, Kramer SJ, Cesarin BJ, Schooley DA. Isolation and primary structure of the eclosion hormone of the tobacco hornworm, Manduca sexta. Biochem Biophys Res Commun 1987; 146: 746 –750.
35.
Kingan TG, Adams ME. Ecdysteroids regulate secretory competence in Inka cells. J Exp Biol 2000; 203: 3011–3018.
36.
Kingan TG, Gray W, Zitnan D, Adams ME. Regulation of ecdysis-triggering hormone release by eclosion hormone. J Exp Biol 1997; 200: 3245–3256.
37.
Klein C, Kallenborn HG, Radlicki C. The “Inka cell” and its associated cells: Ultrastructure of the epitracheal glands in the gypsy moth, Lymantria dispar. J Insect Physiol 1999; 45: 65–73.
38.
Klukas KA, Brelje TC, Mesce KA. Novel mouse IgG-like immunoreactivity expressed by neurons in the moth Manduca sexta: Developmental regulation and colocalization with crustacean cardioactive peptide. J Microsc Res Tech 1996; 35: 242–264.
39.
Kostron B, Kaltenhauser U, Seibel B, Braunig P, Honegger HW. Localization of bursicon in CCAP-immunoreactive cells in the thoracic ganglia of the cricket Gryllus bimaculatus. J Exp Biol 1996; 199: 367–377.
40.
Kostron B, Market D, Kellermann J, Carter CE, Honegger HW. Antisera against Periplaneta americana Cu,Zn-superoxide dismutase (SOD): Separation of the neurohormone bursicon
HORMONES AND INSECT MOLTING
197
from SOD, and immunodetection of SOD in the central nervous system. Insect Biochem Mol Biol 1999; 29(10): 861– 871. 41.
Lehman HK, Murgiuc CM, Miller TA, Lee TD, Hildebrand JG. Crustacean cardioactive peptide in the sphinx moth, Manduca sexta. Peptides 1993; 14: 735–741.
42.
Levine RB, Truman JW. Dendritic reorganization of abdominal motoneurons during metamorphosis of the moth, Manduca sexta. J Neurosci 1985; 5: 2424 –2431.
43.
Loi PK, Emmal SA, Park Y, Tublitz NJ. Identification, sequence and expression of a crustacean cardioactive peptide (CCAP) gene in the moth Manduca sexta. J Exp Biol 2001; 204: 2803–2816.
44.
Marder E, Bucher D. Central pattern generators and the control of rhythmic movements. Curr Biol 2001; 11: R986 –R996.
45.
Marti T, Takio K, Walsh KA, Terzi G, Truman JW. Microanalysis of the amino acid sequence of the eclosion hormone from the tobacco hornworm Manduca sexta. FEBS Lett 1987; 219: 415– 418.
46.
McNabb SL, Baker JD, Agapite J, Steller H, Riddiford LM, Truman JW. Disruption of a behavioral sequence by targeted death of peptidergic neurons in Drosophila. Neuron 1997; 19: 813– 823.
47.
McNeil GP, Zhang X, Genova G, Jackson FR. A molecular rhythm mediating circadian clock output in Drosophila. Neuron 1998; 20(2): 297–303.
48.
Mesce KA, Klukas KA, Brelje TC. Improvements for the anatomical characterization of insect neurons in whole mount: The use of cyanine-derived fluorophores and laser scanning confocal microscopy. Cell Tissue Res 1993; 271: 381–397.
49.
Mesce KA, Truman JW. Metamorphosis of the ecdysis motor pattern in the hawkmoth, Manduca sexta. J Comp Physiol A 1988; 163: 287–299.
50.
Miles CI, Weeks JC. Developmental attenuation of the larval pre-ecdysis motor pattern in the tobacco hornworm, Manduca sexta. J Comp Physiol A 1991; 168(2): 179 –190.
51.
Miles CI, Weeks JC. Developmental attenuation of the larval pre-ecydsis motor pattern in the tobacco hornworm, Manduca sexta. J Comp Physiol A 1991; 168: 179 –190.
52.
Morton DB, Simpson PJ. Cellular signaling in eclosion hormone action. Journal of Insect Physiology 2002; 42(1): 1–13.
53.
Nassel DR. Functional roles of neuropeptides in the insect central nervous system. Naturwissenschaften 2000; 87(10): 439 – 449.
54.
Naya S, Suzuki K, Fugo H, Sehnal F. Eclosion hormone-like immunoreactivity in the nervous system of Bombyx mori (Lepidoptera, Bombycidae) and Antheraea yamamai (Lepidoptera, Saturnidae) before and after hatching. European J Entomol 1994; 91: 189 –196.
55.
Nijhout HF. Insect Hormones. Princeton, NJ: Princeton University Press, 1994.
56.
Novicki A, Weeks JC. Organization of the larval pre-ecdysis motor pattern in the tobacco hornworm, Manduca sexta. J Comp Physiol A 1993; 173: 151–162.
57.
Novicki A, Weeks JC. The initiation of pre-ecdysis and ecdysis behaviors in larval Manduca sexta—The roles of the brain, terminal ganglion, and eclosion hormone. J Exp Biol 1996; 199: 1757–1769.
58.
Novicki A, Weeks JC. The initiation of pre-ecdysis and ecdysis behaviors in larval Manduca sexta: The roles of the brain, terminal ganglion and eclosion hormone. J Exp Biol 1996; 199(8): 1757–1769.
59.
O’Brien MA, Taghert PH. A peritracheal neuropeptide system in insects—Release of myomodulin-like peptides at ecdysis. J Exp Biol 1998; 201: 193–209.
60.
Orchard I, Lange AB, Bendena WG. FMRFamide-related peptides: A multifunctional family of structurally related neuropeptides in insects. Advances in Insect Physiology. 2001; 28: 267–329.
198
MESCE AND FAHRBACH
61.
Park Y, Zitnan D, Gill SS, Adams ME. Molecular cloning and biological activity of ecdysistriggering hormones in Drosophila melanogaster. FEBS Lett 1999; 463(1–2): 133–138.
62.
Phlippen MK, Webster SG, Chung JS, Dircksen H. Ecdysis of decapod crustaceans is associated with a dramatic release of crustacean cardioactive peptide into the haemolymph. J Exp Biol 2000; 203(3): 521–536.
63.
Predel R, Eckert M. Neurosecretion: Peptidergic systems in insects. Naturwissenschaften 2000; 87(8): 343–350.
64.
Reynolds SE, Truman JW. Eclosion hormone and bursicon titres and the onset of hormonal responsiveness during the last day of adult development in Manduca sexta (L.). J Exp Biol 1979; 78: 77– 86.
65.
Riddiford LM, Hewes RS, Truman JW. Dynamics and metamorphosis of an identifiable peptidergic neuron in an insect. J Neurobiol 1994; 25: 819 – 830.
66.
Schachtner J, Klaassen L, Truman JW. Metamorphic control of cyclic guanosine monophosphate expression in the nervous system of the tobacco hornworm, Manduca sexta. J Comp Neurol 1998; 396: 238 –252.
67.
Stangier J, Hilbich C, Dircksen H, Keller R. Distribution of a novel cardioactive neuropeptide (CCAP) in the nervous system of the shore crab Carcinus maenas. Peptides 1988; 9: 795– 800.
68.
Swensen AM, Marder E. Modulators with convergent cellular actions elicit distinct circuit outputs. J Neurosci 2001; 21: 4050 – 4058.
69.
Trube A, Audehm U, Dircksen H. Crustacean cardioactive peptide-immunoreactive neurons in the ventral nervous system of crayfish. J Comp Neurol 1994; 348: 80 –93.
70.
Truman JW. Physiology of insect ecdysis. I. The eclosion behaviour of Saturniid moths and its hormonal release. J Exp Biol 1971; 54: 805– 814.
71.
Truman JW. Physiology of insect ecdysis. III. The relationship between the hormonal control of eclosion and of tanning in the tobacco hornworm, Manduca sexta. J Exp Biol 1973; 58: 821– 829.
72.
Truman JW. The preparatory behavior rhythm of the moth Manduca sexta: An ecdysteroidtriggered circadian rhythm that is independent of the brain. J Comp Physiol A 1984; 155: 521–528.
73.
Truman JW. The eclosion hormone system of insects. Prog Brain Res 1992; 92: 361–374.
74.
Truman JW. Ecdysis control sheds another layer. Science 1996; 271: 40 – 41.
75.
Truman JW, Copenhaver PF. The larval eclosion hormone neurones in Manduca sexta: Identification of the brain-proctodeal neurosecretory system. J Exp Biol 1989; 147: 457– 470.
76.
Truman JW, Riddiford LM. Neuroendocrine control of ecdysis in silkmoths. Science 1970; 167: 1624 –1626.
77.
Truman JW, Taghert PH, Copenhaver PF, Tublitz NJ, Schwartz LM. Eclosion hormone may control all ecdyses in insects. Nature 1981; 291: 70 –71.
78.
Turbeville JM, Pfeifer DM, Field KG, Raff RA. The phylogenetic status of arthropods, as inferred from 18S rRNA sequences. Mol Biol Evol 1991; 8: 669 – 686.
79.
Weeks JC, Truman JW. Neural organization of peptide-activated ecdysis behaviors during the metamorphosis of Manduca sexta. II. Retention of the proleg motor pattern despite loss of the prolegs at pupation. J Comp Physiol A 1984; 155: 423– 433.
80.
Williams JA, Su HS, Bernards A, Field J, Sehgal A. A circadian output in Drosophila mediated by neurofibromatosis-1 and Ras/MAPK. Science 2001; 293(5538): 2251–2256.
81.
Zhang X, McNeil GP, Hilderbrand-Chae MJ, Franklin TM, Schroeder AJ, Jackson FR. Circadian regulation of the lark RNA-binding protein within identifiable neurosecretory cells. J Neurobiol 2000; 45(1): 14 –29.
82.
Zitnan D, Adams ME. Excitatory and inhibitory roles of central ganglia in initiation of the insect ecdysis behavioural sequence. J Exp Biol 2000; 203: 1329 –1340.
HORMONES AND INSECT MOLTING 83.
199
Zitnan D, Kingan TG, Hermesman JL, Adams ME. Identification of ecdysis-triggering hormone from an epitracheal endocrine system. Science 1996; 271: 88 –91. 84. Zitnan D, Ross LS, Zitnanova I, Hermesman JL, Gill SS, Adams ME. Steroid induction of a peptide hormone gene leads to orchestration of a defined behavioral sequence. Neuron 1999; 23: 523–535. 85. Zitnanova I, Adams ME, Zitnan D. Dual ecdysteroid action on the epitracheal glands and central nervous system preceding ecdysis of Manduca sexta. J Exp Biol 2001; 204: 3483– 3495.
Integration of Endocrine Signals That Regulate Insect Ecdysis Karen A. Mesce* and Susan E. Fahrbach† *Departments of Entomology and Neuroscience and Graduate Program in Neuroscience, University of Minnesota, St. Paul, Minnesota 55108; and †Department of Entomology and Neuroscience Program, University of Illinois at Urbana–Champaign, Urbana, Illinois 61801
The extremely large number of insects and members of allied groups alive today suggests that molting—shedding of an old cuticle—may be one of the most commonly performed behaviors on our planet. Removal of an old cuticle in insects is associated with stereotyped, species-specific patterns of behavior referred to as ecdysis. It has been recognized for decades that the initiation of ecdysis is under hormonal control, but until recently many of the key peptides that regulate ecdysis were unknown. The report in 1996 of a new ecdysis-triggering hormone (ETH) sparked an era of significant advances in our understanding of the regulation of molting. This article summarizes the current model of peptide regulation of ecdysis, a model that is based on a positive feedback loop between ETH and a brain peptide, eclosion hormone. Then the relationship of these regulatory peptides to the neural circuitry that is the ultimate driver of the behavior are described. Because insects can undergo both status quo (larval–larval) and metamorphic (larval–pupal and pupal–adult) molts, differences in ecdysis behavior at different life stages are described and potential sources of these differences are identified. Most of the work described is based on studies of ecdysis in the hawkmoth, Manduca sexta, but results from studies of ecdysis in the fruit fly Drosophila melanogaster are also discussed. KEY WORDS: bursicon; CCAP; ecdysis; ecdysis-triggering hormone; eclosion; eclosion hormone; epitracheal glands; Inka cell; Manduca sexta; molting; VM neuron. ©
2002 Elsevier Science (USA)
INTRODUCTION
Insect species appear in the fossil record approximately 425 million years ago and today abound by the millions, a testimonial to their differential reproductive success. In part, this success, which reflects an extraordinary ability to adapt to varying environmental conditions, can be attributed to their sophisticated endocrine systems. The rich behavioral repertoires expressed by insects owe their flexibility and operational range to an overlay of endocrine factors that expands the capabilities of the relatively simple arthropod nervous system. All aspects of an insect’s life are intimately associated with its endocrinology. Chief among the insect hormones are the ecdysteroids (steroids), juvenile Address correspondence and reprint requests to S. E. Fahrbach, Department of Entomology, University of Illinois at Urbana–Champaign, 320 Morrill Hall, 505 S. Goodwin Avenue, Urbana, IL 61801. Fax: (217) 244-3499. E-mail: [email protected]. 179 ©
0091-3022/02 $35.00 2002 Elsevier Science (USA) All rights reserved.
180
MESCE AND FAHRBACH
hormones (terpinoids), peptides, and biogenic amines (55). Recent reviews have examined the actions of neuropeptides on visceral tissues, muscle, and the central and peripheral nervous systems (4, 53, 60, 63). The biological functions of octopamine, a key neuromodulator-neurohormone in insects, have also been recently reviewed (3) and therefore are not considered here. Other recent reviews have considered the ecdysteroids as mediators of neural plasticity (19) and provide an historical perspective on our understanding of the neuropeptide regulation of ecdysis behavior (14). The present review has more narrow goals. One is to describe how a remarkably elaborate cascade of endocrine factors regulates the seemingly simple behavioral task of shedding the cuticle in a way we believe will be accessible to the nonspecialist. A second goal is to compare what is known about the control of ecdysis behavior in larvae, pupae, and adults, the three major life stages of insects with complete metamorphosis. Our aim is to integrate a series of recent studies and unpublished results to illustrate the intricate neuroendocrine relationships required for expression of ecdysis-related behaviors. As one might expect, the study of insect molting requires investigation at both organismal and cellular–molecular levels. Physiological and behavioral studies have traditionally used large insects to study ecdysis, particularly the tobacco hornworm Manduca sexta (18, 70, 76). Such studies are now profitably being extended with molecular genetic studies of the fruit fly Drosophila melanogaster, the first postgenomic insect. Because progress in our understanding of the hormonal regulation of ecdysis cascades is currently rapid, this review augments other recent reviews on the topic (25, 29, 52, 73).
HISTORICAL PERSPECTIVE AND OVERVIEW OF ECDYSIS
This is still an era of discovery for insect hormones. The number of hormones known to be involved in the regulation of ecdysis-related motor programs has grown significantly over the past 5 years. Because the accompanying nomenclature is potentially confusing, careful attention is given to the definition of key terms used in this article. At the end of each molt cycle (the production of a new cuticle by the monolayer of epidermal cells that produce the insect exoskeleton), insects perform stereotyped, species-specific behaviors to shed the cuticle of the previous stage in a process termed “ecdysis”. A separate term, eclosion, is typically reserved for the final ecdysis of the insect’s life, so that adult ecdysis is often referred to as adult eclosion or simply eclosion. Over 30 years ago, blood-borne eclosion hormone (EH) 1 activity was first defined on the basis of its ability to 1 Abbreviations used: 20-E, 20-hydroxyecdysone; A1, A2 etc., abdominal ganglia of the ventral nerve cord; CCAP, crustacean cardioactive peptide; cGMP, cyclic 3⬘,5⬘-guanosine monophosphate; CNS, central nervous system; DL, dorsal longitudinal muscle; EH, eclosion hormone; ETH, ecdysis-triggering hormone; LDI, long distance interneuron; MN-2, motor neuron 2 of Manduca abdominal ganglia; MN-3. motor neuron 3 of Manduca abdominal ganglia; PETH, pre-ecdysis-
HORMONES AND INSECT MOLTING
181
initiate ecdysis behavior in pharate adult giant silkmoths (70, 76). (The term “pharate” refers to an insect fully developed and ready to molt, but still encased in the cuticle of the preceding stage.) Subsequent studies in the now more commonly used North American hawkmoth, Manduca sexta (hereafter referred to as Manduca), have revealed that EH also elicits ecdysis behavior when the insect molts from one larval stage to the next (termed larval ecdysis) and from the larval to the pupal stage (pupal ecdysis). Eclosion hormone is now identified as a 62-amino-acid peptide synthesized exclusively in two pairs of ventral medial (VM) neurons located in the protocerebral region of the brain (26, 32–34, 45, 54, 75). Although Manduca is the primary species used for research into the effects of EH on behavior, EH has also been shown to be synthesized by VM neurons and to initiate ecdysis behavior in nonlepidopteran insect species, including Drosophila (30, 73, 77). Eclosion hormone is currently assumed to regulate all insect ecdyses. The central nervous system (CNS) of insects consists of a brain and a chain of segmental neural ganglia, referred to as the ventral nerve cord (5). The motor neurons that drive the motor patterns of ecdysis reside in the segmental ganglia of the abdomen and the thorax; their axons innervate the musculature of the exoskeleton via segmentally repeated dorsal and ventral nerves (42). The life cycle of Manduca consists of five larval stages, a pupal stage during which adult development occurs, and the adult (the life cycle of Drosophila is similar, although there are only three larval stages in this species.) Each stage is separated by an episode of ecdysis in which the cuticle of the former stage is shed. In the case of Manduca and many other insects, the performance of ecdysis behavior removes the old cuticle by a series of coordinated abdominal movements characterized by an element of an anteriorly directed peristalsis. Adult eclosion in Manduca, however, differs from both larval and pupal ecdysis in one highly significant aspect: No consistent pattern of preecdysis rhythmic motor activity is expressed as a prelude to adult eclosion. In sharp contrast to pharate larvae and pupae, the pharate adult remains fairly quiescent except for occasional bouts of body twitching (Fig. 1A). Prior to each molt, hemolymph titers of ecdysteroids (primarily 20-hydroxyecdysone, or 20-E) rise and then return to basal levels (55); without the decline, ecdysis is blocked, and recent studies indicate that both central and peripheral actions of the steroid are important in preparing the insect for each molt (35, 84, 85). The decline in circulating steroids likely acts in concert with other signals that either specify the insect’s readiness to molt or provide circadian cues so that the insect molts during the appropriate phase of the light– dark cycle (74).
triggering hormone; SEG, subesophageal ganglion; T1-T3, the prothoracic, mesothoracic, and metathoracic ganglia of the ventral nerve cord; TSM, tergosternal muscle; TX, thorax; VM, ventral medial neurons of the insect brain that synthesize and secrete EH.
182
MESCE AND FAHRBACH
FIG. 1. Extracellular recordings from the tergosternal muscles (TSMs) in the fourth through sixth abdominal segments (A4 –A6) and from the dorsal longitudinal (DL) muscles in the thorax (TX) of an intact pharate adult Manduca. Muscle activity was monitored beginning approximately 5– 6 h before the predicted time of adult ecdysis to determine if any pre-ecdysis-like motor patterns were present. Electrophysiological methods used are as previously described (49). (A) Records of muscle activity expressed throughout various time periods prior to eclosion are shown; samples of activity patterns presented were randomly chosen. In over 40 insects examined in this way, no preecdysis motor pattern was observed. Activity in the thorax was minimal and remained unpatterned until eclosion. Intersegmental TSM coactivity was always variable, of short duration (reflecting a rapid body twitching), and intermittent. (B) At the onset of eclosion behavior, the DL muscles first showed strong activity, which was always coactive with TSM excitation and correlated with synchronous lateral body contractions that caused the abdomen to extend. By the time of the sixth extension cycle, the peristalsis component appeared (marked by asterisks) and continued, while involving more posterior segments (not shown). This rapid anteriorly directed peristalsis caused the retraction of the abdomen, which was followed by the abdominal extension, a pattern that was repeated for many cycles. The temporal parameters of these extension bouts were indistinguishable from the activity patterns expressed during larval and pupal preecdysis behavior.
HORMONES AND INSECT MOLTING
183
FIG. 2. (A) Schematic diagram of the peripheral peritracheal neuroendocrine system of insects. (B) Summary diagram of the hormonal and cellular interactions leading to events involved in ecdysis-related behaviors derived from research on Manduca and Drosophila. Boxes contain both the names of ecdysis-related hormones (or factors) and the specific cells that secrete them. Hatched lines indicate relationships that are suggested by current research and for which comparatively less is known.
PREECDYSIS BEHAVIOR AND ITS ENDOCRINE CONTROL
The distinctive preecdysis motor patterns observed in larval and prepupal insects loosen the attachments between the new and the old cuticle prior to ecdysis. In Manduca larvae and prepupae, two distinct phases of preecdysis behavior, designated preecdysis I and preecdysis II, have been identified (84). These preecdysis behaviors are regulated by peptide hormones secreted by 18 segmentally repeated epitracheal glands located near the spiracular tracheae of the abdominal segments (Fig. 2A). The epitracheal glands, which were first described in 1996 (37, 83), each consist of a small number of cells (four per gland in Manduca), one of which is referred to as the Inka cell. Inka cells are characterized by synthesis and secretion of a myomodulin-like peptide (59) and
184
MESCE AND FAHRBACH
at least three additional peptides, two of which, preecdysis-triggering hormone (PETH) and ecdysis-triggering hormone (ETH), activate preecdysis I and preecdysis II (83, 84). The myomodulin-like peptide is released from the Inka cells of Drosophila and Manduca in the hour prior to all ecdyses and is present in putative Inka cell homologs in various insects such as beetle, dragonfly, and mosquito, but at present nothing is reported concerning its ecdysis-related functions (59). PETH and ETH were first described in Manduca and have been subsequently identified in another lepidopteran, the silkmoth, Bombyx mori (1), and in Drosophila (61). Both peptides are encoded by a single gene downstream of a promoter region that contains ecdysone response elements (84). In the Inka cells, a precursor peptide is cleaved to produce the 11-amino-acid PETH, the 26-amino-acid ETH, and a 46-amino-acid peptide of unknown function referred to as ETH-associated peptide (ETH-AP). Application of PETH to the isolated CNS of Manduca can activate a known neural correlate of preecdysis I behavior, strong bursts of motor activity in the dorsal nerves. This pattern of electrical activity in the CNS is often referred to as the fictive component of preecdysis behavior. The bursts in the dorsal nerves are in phase with the synchronous dorsoventral compressions of the insect’s abdomen that occur during the first 20 or so min of preecdysis behavior (51, 56, 57, 84). Animals injected only with PETH express preecdysis I behavior exclusively, and no other behaviors associated with the ecdysis sequence such as preecdysis II or ecdysis are subsequently displayed (84). In contrast to the actions of PETH, currently believed to be restricted to controlling preecdysis, ETH also indirectly activates the behavioral routines of ecdysis itself in a chain of events that are described in more detail in following sections (83). The most direct action of this peptide, however, is the initiation of preecdysis II behavior (84). Injections of low physiological concentrations of ETH into pharate larvae previously exposed to PETH result in the selective expression of asynchronous posterior-ventral contractions of abdominal body segments coupled with repeated retraction-extensions of the prolegs. A fictive behavioral correlate of this preecdysis II pattern has also been identified: it consists of strong motor bursts in the ventral nerves in the absence of strong bursting in the dorsal roots (84). Fictive preecdysis II can be elicited with low concentrations of ETH. Application of higher ETH concentrations (100 –300 nM) to isolated ventral nerve cord preparations can elicit ecdysis behavior, with latencies in the range of 22–30 min (84). Under natural conditions, preecdysis I, preecdysis II, and ecdysis occur in sequence. The strong dorsoventral contractions of preecdysis I (roughly a 1-s contraction followed by 3 s of relaxation) are observed first. Preecdysis II behavior in Manduca is then typically observed after the first third of preecdysis I behavior has been performed and continues along with expression of the preecdysis I program until ecdysis is initiated, about 40 min later [see Fig. 6B in reference (84)]. PETH and ETH appear to be co-released from the Inka cells, but a lower response threshold to PETH ensures that preecdysis I is expressed first followed by preecdysis II and then by ecdysis (82, 84).
HORMONES AND INSECT MOLTING
185
ECDYSIS BEHAVIOR AND ITS ENDOCRINE CONTROL
Although the names ascribed to EH and ETH imply that they are the direct hormonal activators of the neural circuits responsible for ecdysis behavior, to date there is no evidence that either hormone is tied to such proximal activation. Instead, it is the neuropeptide crustacean cardioactive peptide (CCAP) that has been shown to have the most direct, shortest latency route to activation of the ecdysis motor program (22). Several studies have indicated that EH stimulates the release of CCAP from neurosecretory neurons distributed along the ventral nerve cord (23, 66). In Manduca, this network consists primarily of segmentally repeated lateral pairs of neurons named cell 27 [or NS-L(neuromere no.)] and interneuron/cell (IN)-704 (7, 12, 38), often referred to jointly as 27/704. These cells were initially identified on the basis of their immunoreactivity to an antibody prepared against CCAP from crustaceans (67) and hence are referred to as CCAP-ir cells. Several lines of evidence indicate that authentic CCAP is present in the insect nervous system. A nonapeptide identical in amino acid sequence to that of CCAP originally isolated from a shore crab, Carcinus maenas, was purified from extracts of the Manduca CNS (6, 41). More recently, the CCAP gene of Manduca was cloned, and it was demonstrated using in situ hybridization that almost all CCAP-ir cells in the CNS express the mRNA coding for this peptide (43). The distribution of CCAP neurons is highly conserved between insects and crustaceans (62, 69). Although extant insect species and crustaccans have evolved as independent lineages for hundreds of millions of years (20, 24, 78), CCAP has been recently shown to be released during the ecdysis of crabs and crayfish (62), suggesting that its behavior-activating functions have also been conserved. Intracellular recordings made from abdominal CCAP neurons in pharate larvae of Manduca revealed that EH rapidly elevated cGMP levels within these cells, which in turn resulted in increased release of CCAP (23). When latencies to ecdysis were compared after application of various peptides to the isolated nervous system, the response to CCAP (5 min) was significantly shorter than the response to either EH (26 min) or ETH (40 min) (23). CCAP can also induce ecdysis at developmentally inappropriate times (i.e., intermolt stages, when the new cuticle is incomplete), periods when the CNS is refractory to ETH and EH action (23). At present, therefore, no peptide is linked more tightly to the onset of ecdysis than CCAP. This makes it particularly unfortunate that no information is currently available about the neuronal targets of CCAP at the level of ecdysis pattern-generating circuitry. Although a number of motor neurons with ecdysis-related patterns of activity have been identified (79), the individual central pattern generator neurons that generate the motor patterns underlying ecdysis behavior have not been identified. This is clearly a challenging area for future research. Figure 2B shows the coordinated hormonal events associated with the expression of ecdysis behavior. To summarize, at the end of the molt cycle, when the new cuticle is formed and a successful molt is possible, the Inka cells of the
186
MESCE AND FAHRBACH
epitracheal glands and the VM neurons of the brain release their respective peptide hormones, PETH/ETH and EH (27, 35, 84). At present it cannot be conclusively stated that either EH or ETH is released first, as each hormone has been clearly demonstrated to cause the release of the other (13, 23, 36). Arguments have been made for the primacy of PETH/ETH based on in vitro differences in the responses to PETH, ETH, and EH (82). What is clear is that reciprocal positive feedback mechanisms ensure the attainment of successively higher levels of PETH/ETH and EH. When sufficiently high levels of circulating ETH are achieved, the VM neurons of the brain respond by producing a surge of EH released both centrally and peripherally into the hemolymph. Such a coordinated release is made possible by the extensive projections of the VM neurons within each segmental ganglion as well as projections to a peripheral neurohemal release site via the proctodeal nerve (75). One of the actions of EH on the CCAP-containing 27/704 cells is to facilitate the release of CCAP (23). CCAP then serves as the proximate trigger of ecdysis behavior.
BURSICON IN THE CRUSTACEAN CARDIOACTIVE PEPTIDE NETWORK
Adding a further level of complexity, a subset of the neurons of the CCAP network also contain a neuropeptide called bursicon. Bursicon was originally defined as the primary factor that promotes the hardening and tanning (sclerotization) of new cuticle after the insect has completed ecdysis (64). New studies indicate that, at a minimum, the CCAP cell 27 in Manduca and its homologs in other insects likely contain bursicon (39, 28). An attempt to determine the complete amino acid sequence of the bursicon hormone is currently underway in the laboratory of H.-W. Honegger (Vanderbilt University, TN). The microsequencing of bursicon hormone purified from cockroaches recently yielded several partial peptide sequences (40). Subsequent immunocytochemical studies using antisera directed against these partial sequences confirmed that the CCAP-ir cell 27 in Manduca and the CCAP-ir cell 27 homologs in other insects, including crickets, cockroaches, and several species of flies, are immunoreactive to these antisera (28). These results suggest that specific bursicon amino acid sequences are conserved even among distantly related insect species. Studies in Manduca have demonstrated an association between EH release and the subsequent secretion of bursicon into the hemolymph, although a full picture is lacking because published reports have focused almost exclusively on adult eclosion (64, 71). The important issue of how the actions of bursicon are temporally restricted, however, remains unresolved. For the Manduca pharate adult, in particular, it is essential that tanning of the cuticle not be activated until the insect has removed its old cuticle, surfaced from its underground pupation site and gained a perch from which it can inflate its wings. The premature action of bursicon before completion of the ecdysis motor program could result in a premature hardening of the cuticle and a blocking of wing inflation that would likely be lethal. The observation that CCAP and
HORMONES AND INSECT MOLTING
187
bursicon are often colocalized in cell 27 therefore provides a neuroendocrine puzzle. Studies are needed to determine whether these hormones are differentially secreted. Whether bursicon is modified in some way to produce its delayed (45– 60 min) action during adult ecdysis or has additional actions that block premature sclerotization are questions that remain unanswered.
ADDITIONAL FACTORS CONTROLLING ECDYSIS
Although strong evidence indicates that EH is the link between ETH and CCAP action (23), a number of intriguing recent studies suggest the presence of additional layers of control, some of which involve descending neural activity (2, 58, 82). Ecdysis-triggering hormone has been shown to excite the EH-containing VM neurons (13) and to elicit a broadening of the VM cell spike (23). Electrical stimulation of these cells causes elevation of cGMP in the CCAP 27/704 neural network. However, when the part of the brain (protocerebrum) that contains the VM neurons was removed from pharate Manduca larvae (the deutocerebrum was also removed, leaving the tritocerebral lobes intact), 10 of 14 insects (71%) injected with ETH expressed ecdysis behavior at the expected time (82). Removal of the subesophageal ganglion (SEG) or thoracic ganglia severely limited or prevented the onset of ecdysis behavior. In turn, application of ETH to isolated nerve cords with the SEG intact but the brain ablated elicited an unmistakable ecdysis motor response and concomitant elevations in cGMP among the CCAP cells. Such results suggest that, although ETH-induced release of EH from the VM neurons is sufficient to activate signaling cascades resulting in CCAP release, this pathway may not be required. Studies in Drosophila (2, 46) further suggest that the VM neurons in the brain may not be required for ecdysis. Genetically modified knockout flies that lack the VM neurons expressed components of adult eclosion behavior and emerged from the pupal cuticle; the behavior of these pharate adults, however, was described as weak and poorly organized. In addition, the flies that did succeed in eclosing could not inflate their wings, possibly as a consequence of abnormal bursicon release (46). In addition, the pharate adult knockout flies lacking the VM neurons were no longer sensitive to ETH. Although ETH has not yet been shown to activate a specific behavior in pharate adult flies, treatment of pharate adult flies with exogenous ETH can advance the onset of adult eclosion (2, 46), and a gene encoding ETH-like peptides has been identified in Drosophila (61). The lack of a response to ETH in the knockout flies suggests that, in wild-type flies, ETH, EH, and/or sensory inputs must converge on some unidentified neural or hormonal element(s) in the CNS in order to drive activity in the CCAP cell 27/704 network. In the absence of the ETH-sensitive VM neurons, ecdysis activation can apparently still be achieved by direct activation of downstream parts of the pathway. Whether these observations on Drosophila can be generalized to other insects is currently unknown.
188
MESCE AND FAHRBACH
In addition, the switch from larval preecdysis to ecdysis behaviors appears not to be a simple consequence of CCAP release. Studies by Zitnan and Adams (82) have demonstrated that an inhibitory influence from the SEG, thoracic ganglia, and first abdominal ganglion must be removed before CCAP-induced ecdysis behavior can be initiated. After initiation of spontaneous or ETHevoked preecdysis behavior, isolation of the abdomen from anterior ganglia (ligation) significantly advanced the onset of ecdysis in all insect tested. Removal of the head has also been shown to induce eclosion onset in pharate adult flies and in Manduca (13). Because ligations that separate the head from the body in pharate adults can elicit eclosion within minutes or even seconds (K. A. Mesce, personal observation), such massive severing of neural connections may result in an injury-induced excitation of stimulatory descending neural signals rather than a simple removal of inhibitory elements. Such a scenario is typically overlooked. A picture emerges in which descending neural control mechanisms are coupled with hormonal feedback loops to ensure that an insect initiates ecdysis only when the insect is physiologically prepared to survive the ecdysis event (the formation of the new cuticle is complete and the digestion of the old cuticle is sufficiently advanced). As previously noted, which trigger has primacy is currently impossible to say and may differ among insect species.
CRUSTACEAN CARDIOACTIVE PEPTIDE NEURONS AND STAGE-SPECIFIC BEHAVIORS
The morphology of the CCAP cells indicates that both central and peripheral release are possible (7, 15, 38). It has been argued that the CCAP-containing neurons active at the time of ecdysis are those that show immunodetectable elevations in cGMP (12, 21). Although other CCAP neurons are present in the CNS, only the cell 27/704 network, consisting of approximately 50 neurons, expresses cGMP at the time of ecdysis. During adult and pupal ecdysis, however, only the CCAP-ir 27s and not the 704s in the abdomen express increased cGMP (16), while at larval ecdysis the abdominal 704s also express increased cGMP. Thus, local spatial differences in the release of CCAP could account for differences in the activation of ecdysis-related motor programs displayed by different life stages. It was originally noted that the entire cell 27/704 network becomes cGMPimmunopositive about the same time. One recent study, however, has shown that in untreated pharate pupae the 27/704 cells of the SEG and thoracic ganglia show weaker but still detectable cGMP immunoreactivity at 6 h before the onset of ecdysis, well before the release of PETH and ETH from the Inka cells. Zitnan and Adams (82) have suggested that this 27/704 cGMP-ir network in the thoracic ganglia could release an hypothetical ecdysiostatic factor responsible for the descending inhibitory influences described in the previous section. These authors, however, based their idea on the misconception that the thoracic 27/704 network in larvae does not contain CCAP, therefore lead-
HORMONES AND INSECT MOLTING
189
ing them to suggest that this network possesses a different neurohormone. Although there are indeed differences among the thoracic and abdominal 27/704 neurons, as described below, the lack of CCAP is not one of them. Confusion over CCAP expression in the thoracic ganglia stems from several sets of independently published studies in which the results of CCAP immunostaining differed. The study by Klukas et al. (38; see their Figs. 8 and 9) found that the SEG 27/704 cells and the thoracic cell 27s are CCAP-ir in larvae and developing adults. By contrast, another study reported no CCAP staining after the larval stages (8), and other authors mentioned but did not show that the thoracic homologs always express at least weak CCAP immunoreactivity (13). The recent identification of the gene for CCAP in Manduca provides clarification of this issue (43). The published report indicates that there is likely only one CCAP gene in this species. The spatial expression pattern of the CCAP gene using in situ hybridization techniques confirms that the CCAP-ir cells in the larval SEG and thoracic ganglia indeed express the CCAP gene. Later, in the developing adult, the thoracic CCAP-ir 27 and 704 cells continue to exhibit robust staining (Fig. 3). A striking observation that indicates the strength of the association of this peptidergic neural network in Manduca with ecdysis is that, after the final performance of ecdysis by the pharate adult, the CCAP network, with the exception of one cell pair, is dismantled by the programmed cell death of the CCAP neurons (17). Curiously, CCAP expression in the thoracic 27/704 cells of Manduca is highly variable even during larval life as compared to that in the abdominal ganglia. This is especially evident in pharate fifth instar larvae. It is unknown whether such variability represents the transient inability of thoracic cells to generate transcript or reflects intermittent cycles of CCAP synthesis. Other intriguing differences between the thoracic and abdominal homologs have also been documented (15, 38). For example, a gentler tissue fixation protocol is required to immunostain the 27/704 cells of the thoracic ganglia with CCAP antiserum, as if the CCAP epitope is more difficult to preserve in this region of the central nervous system. Differences in CCAP fixation requirements may point to underlying differences in the thoracic cells with regard to CCAP content, storage, and release. Also notable are the other developmentally regulated epitopes and proteins that are differentially expressed among the 27/704 thoracic and abdominal homologs. In fifth instar larvae of Manduca, for example, the 27 cells in the SEG and the 27/704 cells in the three thoracic ganglia all express a 27-kDA membrane-associated protein (38); subsequent microsequencing has revealed novel partial amino acid sequences (K. A. Mesce, unpublished observations). At no time during development do any cell types in the CNS other than the 27/704s express this antigen. A similar pattern of antigen expression in the thoracic ganglia was also evident in larvae stained with the monoclonal antibody P2C5 (17). Studies of P2C5 expression revealed that the 27 cells in the abdomen became P2C5-ir during the pupal and adult stages of development, while comparable cells in the thorax were immunopositive at all life stages. Elimination of 20-E fluctuations during the larval– pupal transition by ligation prevented expression of the P2C5 antigen in these
190
MESCE AND FAHRBACH
FIG. 3. The 27/704 neurons (arrow) immunostained with an antiserum against crustacean cardioactive peptide (CCAP) and a secondary antibody conjugated to Cy3. CCAP-immunoreactive neurons were observed in all ganglia, including the prothoracic ganglion (shown here), of a developing adult 2 days prior to eclosion (Day: ⫺2). Immunostaining protocols and antiserum are as previously described (17, 38). Robust CCAP staining of the 27/704 cells was present in pharate adults and then declined after eclosion, when these cells underwent programmed cell death (17). Scale bar ⫽ 100 m.
abdominal cells (S. E. Fahrbach and C.-M. Wang, unpublished observations), suggesting that regulation of the P2C5 antigen is under the influence of this steroid. Future investigations into the identities and functions of the cellular proteins and antigens just discussed may provide useful information about why and how the thoracic CCAP-27/704 cells differ from those in the abdomen. Because both antigens are associated with cells that express cGMP in response to ETH and EH release, the possibility cannot be dismissed that these factors represent hormone receptor elements and associated signaling proteins.
EXPRESSION OF NOVEL MOTOR PATTERNS AT ADULT ECDYSIS
We now wish to address directly the mechanisms that result in the dramatic differences between the ecdysis-related behaviors of the pharate adult Manduca and those of pharate larvae and pupae (Fig 1B). As stated, the most
HORMONES AND INSECT MOLTING
191
conspicuous difference is the absence of preecdysis I and preecdysis II behaviors prior to the expression of ecdysis behavior (Fig. 1A). Instead of loosening its cuticle via a preecdysis motor program, the pharate adult relies on a series of rotary abdominal movements that are displayed about 10 –12 h before ecdysis. This preparatory behavior, so named to distinguish it from the preecdysis behavior of the larva and the prepupa, is apparently not dependent on ETH or PETH, but instead depends on the decline in ecdysteroid titers that precedes ecdysis (72). Such observations raise several questions. Are preecdysis neural circuits lost during the adult development that occurs after pupal ecdysis? What roles do PETH and ETH play during the last ecdysis? One hypothesis is that the preecdysis pattern generator is retained into the pharate adult stage, but is used to create a novel motor component specific to the adult ecdysis rhythm. The essence of this idea is that separate preecdysis and ecdysis neural circuits are activated together to form a conjoint rhythm, in much the same way as has been demonstrated for neural circuits underlying stomatogastric rhythms in crustaceans (9 –11, 44, 68). The pharate adult performs a repeated series of abdominal extensions and retractions that rely, in part, on the activity of MN-2 and MN-3, the same motor neurons activated during preecdysis and ecdysis behaviors in larvae and pupae. The abdominal retraction phase in the adult consists of a rapidly progressing peristalsis that is likely generated by conserved larval–pupal ecdysis circuitry (49). Though not expressed prior to adult ecdysis, if the preecdysis I neural circuit were operational at the time of ecdysis the motor output of this circuit could constitute the rhythmic extension component observed during adult ecdysis. Support for this idea comes from the analysis of MN-2/MN-3 motor bursts that are repeatedly generated during the extension phase of adult ecdysis. This analysis was accomplished by recording from the tergosternal muscles (TSMs) innervated exclusively by the identified motor neurons MN-2/MN-3, cells used earlier in development to generate preecdysis and ecdysis patterns by larvae and pupae (K. A. Mesce, unpublished observations). Remarkably, the temporal aspects of MN-2/MN-3 (TSM) extension bursts are indistinguishable from MN-2/MN-3 bursting activity generated by larvae during preecdysis behavior. Based on the data of Miles and Weeks (50) for preecdysis activity, the mean period between successive MN-2/MN-3 preecdysis bursts was 8.4 ⫾ 1.2 s (N ⫽ 5), not statistically different from the adult TSM bursts (8.24 ⫾ 0.95 s, N ⫽ 5; p ⬍ 0.001, two-tailed t test). The duration of adult TSM bursts was also similar to the duration of MN-2/MN-3 preecdysis bursts (range of 1.6 –5s). In over 40 animals examined, ecdysis behavior was always marked by three to eight cycles of MN-2/MN-3 coactivity in the absence of the rapid peristaltic (retraction) component. This rapid peristalsis eventually emerged and became interspersed between the extension bursts (Fig. 1B). The period between successive TSM extension bursts remained stable throughout eclosion (49). Independent evidence indicates that an artificial exposure of the isolated CNS to ETH and CCAP can result in the simultaneous expression of ecdysis and preecdysis motor bursts (22). Thus, the novel ecdysis behavior expressed by the pharate adult may be a relatively simple consequence of a conserved ETH-
192
MESCE AND FAHRBACH
activated preecdysis circuit activated simultaneously with the CCAP-dependent circuit responsible for ecdysis. One last point is that the original muscle targets of MN-2/MN-3 (tergopleural and anterior lateral external muscles) are replaced during adult development with the TSMs, which consist of a larger mat of dorsoventrally oriented muscles. Thus, although the magnitude of lateral body constrictions is greater in the adult than in the larva or prepupae, this is not necessarily a consequence of motor neuronal rewiring but could rather reflect muscle remodeling during development. The temporal coordination and modulation of the preecdysis and ecdysis circuits are likely dependent on descending neural inputs, as it has been shown that the removal of descending activity below A2 causes an immediate switch from the adult-specific ecdysis motor pattern to that of the larval-like one (49). Two segmental homologs have been identified that project uninterrupted from A1 and A2 to the terminal abdominal ganglion (Fig. 3). These long distance interneurons (LDIs), when electrically stimulated, cause coactivity of the abdominal TSMs similar to that observed during larval or pupal preecdysis behavior and during the extension phase of adult ecdysis. Because little is known about the CCAP and ETH neuronal targets responsible for ecdysis pattern generation, these newly identified LDI cells provide a potentially fruitful area for future studies of hormonal targets underlying rhythmic behaviors. In addition, because these LDI neurons are present in larvae (K. A. Mesce, unpublished observations), the LDIs could contribute to the descending neural inhibition of ecdysis that can be removed when the connectives are severed between A1 and A2 (82). Interestingly, when larval abdominal ganglia already exposed to CCAP remained connected to T1 and A1, abdominal ganglia 4 – 6 expressed mixed preecdysis and ecdysis bursts. Only when the abdominal ganglia were isolated from T3 and A1 did all of the remaining ganglia exhibit a strong and pure ecdysis motor rhythm. For comparison in the adult, removal of A1 (and, by necessity, A2, as both are part of a larger fused pterothoracic ganglion in the adult) elicits similar strong, larval-like ecdysis bursts. In the case of the adult such activity reflects the unmasking of a motor pattern never normally expressed in its pure larval-like form. In summary, even after EH and CCAP have been released, descending inhibitory neural influences may coordinate the precise timing of ecdysis initiation at all developmental stages. Specifically, in larvae and pupae, the sequential temporal coordination of preecdysis and ecdysis appears to be influenced by descending activity. In the adult, however, possibly the same descending influences may promote a coactivation of the preecdysis and ecdysis motor programs, resulting in the novel ecdysis behavior of the adult (Fig. 4).
CIRCADIAN REGULATION OF ECLOSION: HORMONAL AND CLOCK-OUTPUT MECHANISMS OF BEHAVIOR
One additional important difference between larval–pupal and adult ecdysis behavior in Manduca and Drosophila is that adult eclosion behavior is under
HORMONES AND INSECT MOLTING
193
FIG. 4. Descending long distance interneurons (LDIs) filled retrogradely with the tracer Neurobiotin. Tissue was subsequently reacted with a Cy5-conjugated streptavidin and imaged with the confocal microscope as previously described (48). Insects used were 2 days away from adult ecdysis (Day: ⫺2). (A) Staining of two prominent neurons whose axons in the connectives were filled with tracer after axons were transected below abdominal ganglion 4 (A4). Stained neurons in the first and second thoracic ganglia (T1 and T2) were absent, but were present in the subesophageal ganglion (SEG) (not shown). Neuronal staining in this far anterior ganglion indicated that the lack of filled cells was not the inability of the tracer to reach the thoracic neuromeres. (B) Same Neurobiotin-filled LDIs (arrows) in A1 and A2 of the pterothoracic ganglion. These cells were filled at a level just anterior to the terminal abdominal ganglion (i.e., below A5). Neurons projecting from anterior ganglia to the terminal ganglion were stained in the brain, SEG (both not shown), A1, and A2, but not the thoracic ganglia. Scale bars ⫽ 100 m.
194
MESCE AND FAHRBACH
circadian control. Recent reviews (31, 80) have explored the multiple components and cell molecular mechanisms underlying the clock control of eclosion. Although EH release appears to be (65) under circadian control, recent studies support the idea that multiple clock pathways likely regulate the timing of eclosion, especially because insects lacking their EH-containing VM neurons still show a circadian regulation of eclosion (46). In Drosophila, a class of CCAP-ir neurons to which the cell 27 homologs belong contains the protein known as LARK, which is present in the cellular cytoplasm and shows diurnal changes in abundance (47, 81). This RNA-binding protein has been hypothesized to regulate a translation factor that may mediate CCAP synthesis, secretion, or perhaps neuronal excitability. What regulates LARK itself remains a mystery, but could involve the gated release of EH. Understanding clock mechanisms regulating CCAP content and release will surely provide a more complete picture of the neural and endocrine pathways regulating the expression of ecdysis across insect species One of many remaining puzzles is that the CCAP-ir homologs in Drosophila and other higher dipterans do not express an increase in cGMP at the time of ecdysis, although other cell types and neurons do express such EH-dependent cGMP increases during eclosion. Similarly, expression of the mIgG-like antigen, specific to the CCAP/cGMP-ir neurons of Manduca and other insects (discussed earlier in this article), has never been observed on the CCAP-ir neurons of Drosophila (K. A. Mesce, unpublished observation). Such results indicate that the cGMP-signaling pathways, so highly conserved in the majority of insect orders and through development, may have become modified during evolution of the cyclorrhaphous insects. This alteration likely relates to the unique forms of ecdysis behavior displayed by flies. Possibly, understanding the regulation of LARK may provide clues as to why the cGMP-signaling pathway is absent or reduced in amplitude in the CCAP cells of higher dipterans. FUTURE STUDIES
The diverse array of arthropod species and their disparate environmental constraints provide the comparative neuroendocrinologist with an opportunity both to understand how neuroendocrine control mechanisms become constructed through evolution and how they shift throughout the life of an individual. Yet to date our understanding is based on the study of a very small number of insect species. The greatest opportunities for insect neuroendocrinology in the 21st century will come from a broadening of our perspective beyond the limited number of species currently studied. As ever, shedding of old questions will result in the emergence of new ones for us to solve. ACKNOWLEDGMENTS The authors acknowledge the assistance of Kathleen Klukas in the preparation of the figures. J. Whitfield provided helpful comments on the phylogentic relationship of insects and crustaceans.
HORMONES AND INSECT MOLTING
195
REFERENCES 1.
Adams ME, Zitnan D. Identification of ecdysis-triggering hormone in the silkworm Bombyx mori. Biochem Biophys Res Comm 1997; 230: 188 –191.
2.
Baker JD, McNabb SL, Truman JW. The hormonal coordination of behavior and physiology at adult ecdysis in Drosophila melanogaster. J Exp Biol 1999; 202: 3037–3048.
3.
Braunig P, Pfluger HJ. The unpaired median neurons of insects. Adv Insect Physiol 2001; 28(28): 185–266.
4.
Broeck JV. Neuropeptides and their precursors in the fruitfly, Drosophila melanogaster. Peptides 2001; 22(2): 241–254.
5.
Burrows M. The Neurobiology of an Insect Brain. Oxford, UK: Oxford Univ. Press, 1996.
6.
Cheung CC, Loi PK, Sylwester AW, Lee TD, Tublitz NJ. Primary structure of a cardioactive neuropeptide from the tobacco hawkmoth, Manduca sexta. FEBS Lett 1992; 313: 165–168.
7.
Davis NT, Homberg U, Dircksen H, Levine RB. Crustacean cardioactive peptide immunoreactive neurons in the hawkmoth, Manduca sexta and changes in their immunoreactivity during postembryonic development. J Comp Neurol 1993; 338: 612– 627.
8.
Davis NT, Homberg U, Dircksen H, Levine RB, Hildebrand JG. Crustacean cardioactive peptide-immunoreactive neurons in the hawkmoth Manduca sexta and changes in their immunoreactivity during postembryonic development. J Comp Neurol 1993; 338(4): 612– 627.
9.
Dickinson PS. Interactions among neural networks for behavior. Curr Opin Neurobiol 1995; 5: 792–798.
10.
Dickinson PS, Fairfield WP, Hetling JR, Hauptman J. Neurotransmitter interactions in the stomatogastric system of the spiny lobster: One peptide alters the response of a central pattern generator to a second peptide. J Neurophysiol 1997; 77: 599 – 610.
11.
Dickinson PS, Mecsas C, Marder E. Neuropeptide fusion of two motor-pattern generator circuits. Nature 1990; 344: 155–158.
12.
Ewer J, De Vente J, Truman JW. Neuropeptide induction of cyclic GMP increases in the insect CNS: Resolution at the level of single identifiable neurons. J Neurosci 1994; 14(12): 7704 –7712.
13.
Ewer J, Gammie SC, Truman JW. Control of insect ecdysis by a positive-feedback endocrine system: Roles of eclosion hormone and ecdysis triggering hormone. J Exp Biol 1997; 200(5): 869 – 881.
14.
Ewer J, Reynolds SE. Neuropeptide control of molting in insects. In: Pfaff DW, Arnold A, Etgen A, Fahrbach SE, Rubin R, Eds. Hormones, Brain, and Behavior. San Diego, CA: Academic Press, in press.
15.
Ewer J, Truman JW. Increases in cyclic 3⬘,5⬘-guanosine monophosphate (cGMP) occur at ecdysis in an evolutionarily conserved crustacean cardioactive peptide-immunoreactive insect neuronal network. J Comp Neurol 1996; 370: 330 –341.
16.
Ewer J, Truman JW. Invariant association of ecdysis with increases in cyclic 3⬘,5⬘-guanosine monophosphate immunoreactivity in a small network of peptidergic neurons in the hornworm, Manduca sexta. J Comp Physiol A 1997; 181(4): 319 –330.
17.
Ewer J, Wang CM, Klukas KA, Mesce KA, Truman JW, Fahrbach SE. Programmed cell death of identified peptidergic neurons involved in ecdysis behavior in the moth, Manduca sexta. J Neurobiol 1998; 37: 265–280.
18.
Fahrbach SE. The regulation of neuronal death during insect metamorphosis. BioSci 1997; 47: 77– 85.
19.
Fahrbach SE, Weeks JC. Hormonal regulation of neural and behavioral plasticity in insects. In: Pfaff DW, Arnold A, Etgen A, Fahrbach SE, Rubin R, Eds. Hormones, Brain and Behavior. San Diego, CA: Academic Press, in press.
196
MESCE AND FAHRBACH
20.
Friedrich M, Tautz D. Ribosomal DNA phylogeny of the major extant arthropod classes and the evolution of myriapods. Nature 1995; 376: 165–167.
21.
Gammie SC, Truman JW. An endogenous elevation of cGMP increases the excitability of identified insect neurosecretory cells. J Comp Physiol A 1997; 180: 329 –337.
22.
Gammie SC, Truman JW. Neuropeptide hierarchies and the activation of sequential motor behaviors in the hawkmoth, Manduca sexta. J Neurosci 1997; 17: 4389 – 4397.
23.
Gammie SC, Truman JW. Eclosion hormone provides a link between ecdysis-triggering hormone and crustacean cardioactive peptide in the neuroendocrine cascade that controls ecdysis behavior. J Exp Biol 1999; 202: 343–352.
24.
Giribet G, Ribera C. A review of arthropod phylogeny: New data based on ribosomal DNA sequences and direct optimization. Cladistics 2000; 16: 204 –231.
25.
Hesterlee S, Morton DB. Insect physiology: The emerging story of ecdysis. Curr Biol 1996; 6: 648 – 650.
26.
Hewes RS, Truman JW. TYhe roles of central and peripheral eclosion hormone release in the control of ecdysis behavior in Manduca sexta. J Comp Physiol 1991; 168A: 697–707.
27.
Hewes RS, Truman JW. Steroid regulation of excitability in identified insect neurosecretory cells. J Neurosci 1994; 14(3/2): 1812–1819.
28.
Honegger H-W, Market D, Kostron B, Wilson M, Pierce L, Choi D, Mesce KA. Cellular localization of bursicon using antisera against a partial sequence of this insect cuticlesclerotizing neurohormone. J Comp Neurol, in press.
29.
Horodyski F. Neuroendocrine control of insect ecdysis by eclosion hormone. J Insect Physiol 1996; 42: 917–924.
30.
Horodyski FM, Ewer J, Riddiford LM, Truman JW. Isolation, characterization, and expression of the eclosion hormone gene of Drosophila melanogaster. Eur J Biochem 1993; 215: 221–228.
31.
Jackson FR, Schroeder AJ, Roberts MA, McNeil GP, Kume K, Akten B. Cellular and molecular mechanisms of circadian control in insects. J Insect Physiol 2001; 47: 833– 842.
32.
Kamito T, Tanaka H, Sato B, Nagasawa H, Suzuki A. Nucleotide sequence of cDNA for the eclosion hormnone of the silkworm, Bombyx mori, and the expression in a brain. Biochem Biophys Res Commun 1992; 31: 514 –519.
33.
Kataoka H, Li JP, Lui AS, Kramer SJ, Schooley DA. Complete structure of eclosion hormone of Manduca sexta. Assignment of disulfide bond location. Int J Pept Protein Res 1992; 39: 29 –35.
34.
Kataoka H, Troetschler RG, Kramer SJ, Cesarin BJ, Schooley DA. Isolation and primary structure of the eclosion hormone of the tobacco hornworm, Manduca sexta. Biochem Biophys Res Commun 1987; 146: 746 –750.
35.
Kingan TG, Adams ME. Ecdysteroids regulate secretory competence in Inka cells. J Exp Biol 2000; 203: 3011–3018.
36.
Kingan TG, Gray W, Zitnan D, Adams ME. Regulation of ecdysis-triggering hormone release by eclosion hormone. J Exp Biol 1997; 200: 3245–3256.
37.
Klein C, Kallenborn HG, Radlicki C. The “Inka cell” and its associated cells: Ultrastructure of the epitracheal glands in the gypsy moth, Lymantria dispar. J Insect Physiol 1999; 45: 65–73.
38.
Klukas KA, Brelje TC, Mesce KA. Novel mouse IgG-like immunoreactivity expressed by neurons in the moth Manduca sexta: Developmental regulation and colocalization with crustacean cardioactive peptide. J Microsc Res Tech 1996; 35: 242–264.
39.
Kostron B, Kaltenhauser U, Seibel B, Braunig P, Honegger HW. Localization of bursicon in CCAP-immunoreactive cells in the thoracic ganglia of the cricket Gryllus bimaculatus. J Exp Biol 1996; 199: 367–377.
40.
Kostron B, Market D, Kellermann J, Carter CE, Honegger HW. Antisera against Periplaneta americana Cu,Zn-superoxide dismutase (SOD): Separation of the neurohormone bursicon
HORMONES AND INSECT MOLTING
197
from SOD, and immunodetection of SOD in the central nervous system. Insect Biochem Mol Biol 1999; 29(10): 861– 871. 41.
Lehman HK, Murgiuc CM, Miller TA, Lee TD, Hildebrand JG. Crustacean cardioactive peptide in the sphinx moth, Manduca sexta. Peptides 1993; 14: 735–741.
42.
Levine RB, Truman JW. Dendritic reorganization of abdominal motoneurons during metamorphosis of the moth, Manduca sexta. J Neurosci 1985; 5: 2424 –2431.
43.
Loi PK, Emmal SA, Park Y, Tublitz NJ. Identification, sequence and expression of a crustacean cardioactive peptide (CCAP) gene in the moth Manduca sexta. J Exp Biol 2001; 204: 2803–2816.
44.
Marder E, Bucher D. Central pattern generators and the control of rhythmic movements. Curr Biol 2001; 11: R986 –R996.
45.
Marti T, Takio K, Walsh KA, Terzi G, Truman JW. Microanalysis of the amino acid sequence of the eclosion hormone from the tobacco hornworm Manduca sexta. FEBS Lett 1987; 219: 415– 418.
46.
McNabb SL, Baker JD, Agapite J, Steller H, Riddiford LM, Truman JW. Disruption of a behavioral sequence by targeted death of peptidergic neurons in Drosophila. Neuron 1997; 19: 813– 823.
47.
McNeil GP, Zhang X, Genova G, Jackson FR. A molecular rhythm mediating circadian clock output in Drosophila. Neuron 1998; 20(2): 297–303.
48.
Mesce KA, Klukas KA, Brelje TC. Improvements for the anatomical characterization of insect neurons in whole mount: The use of cyanine-derived fluorophores and laser scanning confocal microscopy. Cell Tissue Res 1993; 271: 381–397.
49.
Mesce KA, Truman JW. Metamorphosis of the ecdysis motor pattern in the hawkmoth, Manduca sexta. J Comp Physiol A 1988; 163: 287–299.
50.
Miles CI, Weeks JC. Developmental attenuation of the larval pre-ecdysis motor pattern in the tobacco hornworm, Manduca sexta. J Comp Physiol A 1991; 168(2): 179 –190.
51.
Miles CI, Weeks JC. Developmental attenuation of the larval pre-ecydsis motor pattern in the tobacco hornworm, Manduca sexta. J Comp Physiol A 1991; 168: 179 –190.
52.
Morton DB, Simpson PJ. Cellular signaling in eclosion hormone action. Journal of Insect Physiology 2002; 42(1): 1–13.
53.
Nassel DR. Functional roles of neuropeptides in the insect central nervous system. Naturwissenschaften 2000; 87(10): 439 – 449.
54.
Naya S, Suzuki K, Fugo H, Sehnal F. Eclosion hormone-like immunoreactivity in the nervous system of Bombyx mori (Lepidoptera, Bombycidae) and Antheraea yamamai (Lepidoptera, Saturnidae) before and after hatching. European J Entomol 1994; 91: 189 –196.
55.
Nijhout HF. Insect Hormones. Princeton, NJ: Princeton University Press, 1994.
56.
Novicki A, Weeks JC. Organization of the larval pre-ecdysis motor pattern in the tobacco hornworm, Manduca sexta. J Comp Physiol A 1993; 173: 151–162.
57.
Novicki A, Weeks JC. The initiation of pre-ecdysis and ecdysis behaviors in larval Manduca sexta—The roles of the brain, terminal ganglion, and eclosion hormone. J Exp Biol 1996; 199: 1757–1769.
58.
Novicki A, Weeks JC. The initiation of pre-ecdysis and ecdysis behaviors in larval Manduca sexta: The roles of the brain, terminal ganglion and eclosion hormone. J Exp Biol 1996; 199(8): 1757–1769.
59.
O’Brien MA, Taghert PH. A peritracheal neuropeptide system in insects—Release of myomodulin-like peptides at ecdysis. J Exp Biol 1998; 201: 193–209.
60.
Orchard I, Lange AB, Bendena WG. FMRFamide-related peptides: A multifunctional family of structurally related neuropeptides in insects. Advances in Insect Physiology. 2001; 28: 267–329.
198
MESCE AND FAHRBACH
61.
Park Y, Zitnan D, Gill SS, Adams ME. Molecular cloning and biological activity of ecdysistriggering hormones in Drosophila melanogaster. FEBS Lett 1999; 463(1–2): 133–138.
62.
Phlippen MK, Webster SG, Chung JS, Dircksen H. Ecdysis of decapod crustaceans is associated with a dramatic release of crustacean cardioactive peptide into the haemolymph. J Exp Biol 2000; 203(3): 521–536.
63.
Predel R, Eckert M. Neurosecretion: Peptidergic systems in insects. Naturwissenschaften 2000; 87(8): 343–350.
64.
Reynolds SE, Truman JW. Eclosion hormone and bursicon titres and the onset of hormonal responsiveness during the last day of adult development in Manduca sexta (L.). J Exp Biol 1979; 78: 77– 86.
65.
Riddiford LM, Hewes RS, Truman JW. Dynamics and metamorphosis of an identifiable peptidergic neuron in an insect. J Neurobiol 1994; 25: 819 – 830.
66.
Schachtner J, Klaassen L, Truman JW. Metamorphic control of cyclic guanosine monophosphate expression in the nervous system of the tobacco hornworm, Manduca sexta. J Comp Neurol 1998; 396: 238 –252.
67.
Stangier J, Hilbich C, Dircksen H, Keller R. Distribution of a novel cardioactive neuropeptide (CCAP) in the nervous system of the shore crab Carcinus maenas. Peptides 1988; 9: 795– 800.
68.
Swensen AM, Marder E. Modulators with convergent cellular actions elicit distinct circuit outputs. J Neurosci 2001; 21: 4050 – 4058.
69.
Trube A, Audehm U, Dircksen H. Crustacean cardioactive peptide-immunoreactive neurons in the ventral nervous system of crayfish. J Comp Neurol 1994; 348: 80 –93.
70.
Truman JW. Physiology of insect ecdysis. I. The eclosion behaviour of Saturniid moths and its hormonal release. J Exp Biol 1971; 54: 805– 814.
71.
Truman JW. Physiology of insect ecdysis. III. The relationship between the hormonal control of eclosion and of tanning in the tobacco hornworm, Manduca sexta. J Exp Biol 1973; 58: 821– 829.
72.
Truman JW. The preparatory behavior rhythm of the moth Manduca sexta: An ecdysteroidtriggered circadian rhythm that is independent of the brain. J Comp Physiol A 1984; 155: 521–528.
73.
Truman JW. The eclosion hormone system of insects. Prog Brain Res 1992; 92: 361–374.
74.
Truman JW. Ecdysis control sheds another layer. Science 1996; 271: 40 – 41.
75.
Truman JW, Copenhaver PF. The larval eclosion hormone neurones in Manduca sexta: Identification of the brain-proctodeal neurosecretory system. J Exp Biol 1989; 147: 457– 470.
76.
Truman JW, Riddiford LM. Neuroendocrine control of ecdysis in silkmoths. Science 1970; 167: 1624 –1626.
77.
Truman JW, Taghert PH, Copenhaver PF, Tublitz NJ, Schwartz LM. Eclosion hormone may control all ecdyses in insects. Nature 1981; 291: 70 –71.
78.
Turbeville JM, Pfeifer DM, Field KG, Raff RA. The phylogenetic status of arthropods, as inferred from 18S rRNA sequences. Mol Biol Evol 1991; 8: 669 – 686.
79.
Weeks JC, Truman JW. Neural organization of peptide-activated ecdysis behaviors during the metamorphosis of Manduca sexta. II. Retention of the proleg motor pattern despite loss of the prolegs at pupation. J Comp Physiol A 1984; 155: 423– 433.
80.
Williams JA, Su HS, Bernards A, Field J, Sehgal A. A circadian output in Drosophila mediated by neurofibromatosis-1 and Ras/MAPK. Science 2001; 293(5538): 2251–2256.
81.
Zhang X, McNeil GP, Hilderbrand-Chae MJ, Franklin TM, Schroeder AJ, Jackson FR. Circadian regulation of the lark RNA-binding protein within identifiable neurosecretory cells. J Neurobiol 2000; 45(1): 14 –29.
82.
Zitnan D, Adams ME. Excitatory and inhibitory roles of central ganglia in initiation of the insect ecdysis behavioural sequence. J Exp Biol 2000; 203: 1329 –1340.
HORMONES AND INSECT MOLTING 83.
199
Zitnan D, Kingan TG, Hermesman JL, Adams ME. Identification of ecdysis-triggering hormone from an epitracheal endocrine system. Science 1996; 271: 88 –91. 84. Zitnan D, Ross LS, Zitnanova I, Hermesman JL, Gill SS, Adams ME. Steroid induction of a peptide hormone gene leads to orchestration of a defined behavioral sequence. Neuron 1999; 23: 523–535. 85. Zitnanova I, Adams ME, Zitnan D. Dual ecdysteroid action on the epitracheal glands and central nervous system preceding ecdysis of Manduca sexta. J Exp Biol 2001; 204: 3483– 3495.